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(12) INTERNATIONAL APPLICATION PUBLISHED UNDER THE PATENT COOPERATION TREATY (PCT)

(19) World Intellectual Property Organization

International Bureau

(43) International Publication Date
15 May 2003 (15.05.2003)

PCT

i niri loum n iimi urn im i n ni n in uni mn mil eiiii iiii iinin f m fin an

(10) International Publication Number

WO 03/039483 A2

(51) International Patent Classification 7 : A61K

(21) International Application Number: PCT/US02/36028

(22) International Filing Date:

8 November 2002 (08. 1 1 .2002)

(25) Filing Language:

(26) Publication Language:

English
English

(30) Priority Data:

60/332,349

9 November 2001 (09.1 1.2001) US

(71) Applicant (for all designated States except US): DOW
GLOBAL TECHNOLOGIES INC. [US/US]; Washing-
ton Street, 1790 Building, Midland, MI 48674 (US).

(72) Inventors; and

(75) Inventors/Applicants (for US only): CRANLEY, Paul,
E. [USAJS]; 56 Yaupon Court, Lake Jackson, TX 77566
(US). ALLEN, Jeffrey, R. [US/US]; 14702 Fairtree
Terrace, Poway, CA 92064 (US). DANOWSKI, Kristine,
L. [US/US]; 122 Vail Street, Midland, MI 48642 (US).
MCINTYRE, James, A. [CA/US]; 2115 Burlington
Court, Midland, MI 48642 (US). MILLER, Theodore,
E., Jr. [US/US]; 5902 Woopark, Midland, MI 48640
(US). ROSNER, Bettina, M. [US/US]; 8870 Villa
La Jolla Drive, Apartment 310, La Jolla, CA 92037
(US). STRICKLAND, Alan, D. [US/US]; 115 Hick-
ory, Lake Jackson, TX 77566 (US). SUBRAMANIAN,

Venkiteswaran [US/US]; 3980 Corte Mar De Hierba, San
Diego, CA 92130 (US). SUN, Larry [CA/CA]; 71 Royal
Crescent, Sarnia, Ontario N7S 4Z4 (CA).

(74) Agent: KIMBLE, Karen, L.; The Dow Chemical Com-
pany, Intellectual Property, P.O. Box 1967, Midland, MI
48641-1967 (US).

(81) Designated States (national): AE, AG, AL, AM, AT, AU,
AZ, BA, BB, BG, BR, BY, BZ, CA, CH, CN, CO, CR, CZ,
DE, DK, DM, DZ, EC, EE, ES, FI, GB, GD, GE, GH, GM,
HR, HU, ID, IL, IN, IS, JP, KE, KG, KR, KZ, LC, LK, LR,
LS, LT, LU, LV, MA, MD, MG, MK, MN, MW, MX, MZ,
NO, NZ, OM, PH, PL, PT, RO, RU, SD, SE, SG, SI, SK,
SL, TJ, TM, TO, TR, TT, TZ, UA, UG, US, UZ, YU, ZA,
ZM, ZW.

(84) Designated States (regional): ARIPO patent (GH, GM,
KE, LS, MW, MZ, SD, SL, SZ, TZ, UG, ZM, ZW),
Eurasian patent (AM, AZ, BY, KG, KZ, MD, RU, TJ, TM),
European patent (AT, BE, BG, CH, CY, CZ, DE, DK, EE,
ES, FI, FR, GB, GR, IE, IT, LU, MC, NL, PT, SE, SK,
TR), OAPI patent (BF, BJ, CF, CG, CI, CM, GA, GN, GQ,
GW, ML, MR, NE, SN, TD, TG).

Published:

— without international search report and to be republished .
upon receipt of that report

For two- letter codes and other abbreviations, refer to the "Guid-
ance Notes on Codes and Abbreviations" appearing at the begin-
ning of each regular issue of the PCT Gazette.

(54) Title: AN ENZYME-BASED SYSTEM AND SENSOR FOR MEASURING ACETONE

2e-

electrode

NAD(P) + .

Oxygenase^

NAD(P)H + H + Reductase^ Oxygenase (red) '

H3C-C-CB3OH+ H2O
acetol

H 3 C-C-CK3 + °2

acetone

^ (57) Abstract: Described are enzyme systems specific for acetone and methods of using these enzyme systems to detect acetone in
biological or environmental samples. Biosensors containing these enzyme systems are disclosed, in which detection of acetone may
De achieved by linking electrochemical, photometric, or other detection means to one or more acetone-specific enzyme reactions or
pathways. Methods of using such acetone-specific biosensors include subject management of weight loss, disease detection, and

^ bioavailability monitoring of therapeutics.

WO 03/039483 PCT/US02/36028
AN ENZYME-BASED SYSTEM AND SENSOR FOR MEASURING ACETONE

BACKGROUND OF THE INVENTION

1. Field of the Invention

This invention relates to the field of acetone detection. Acetone-specific enzyme systems
and sensors capable of qualitatively and/or quantitatively detecting acetone have now been
developed, These enzyme systems can be incorporated into relatively inexpensive, simple
and/or portable enzyme-based sensors particularly suited for detecting acetone in
environmental or biological samples, for example, mammalian breath samples.

2. Description of Related Art

A. Acetone Sources

Acetone may be detected in liquids and gases present in or obtained from biological
organisms and various environments. For example, acetone may be detected in
environments such as: natural environments, including soils, sediments, streams, or
wetlands; indoor and outdoor work and home environments; and waste environments,
including waste storage ponds and waste disposal sites. Acetone may be found in
environmental and biological liquids and gases due to introduction of acetone to the
environment or organism from an external source. Thus, environmental acetone may result
from leakage, leaching, waste discharge, or solvent evaporation, or from emission of
combustion gases released by burning wood or plastic or by operating petrochemical
internal combustion engines. Likewise, in a living organism, acetone may be present due to
ingestion, inhalation, or absorption from an external source.

Acetone may also be found in such environmental and biological liquids and gases due to

internal generation of acetone by the environment tested (whether by chemical reaction or

by biological production) or by the organism tested (for example, microbes, animals, etc.).

Thus, the published literature reports that, acetone:

occurs as a biodegradation product of sewage, solid wastes and alcohols, and as an
oxidation product of humic substances. Acetone has been detected in a variety of
plants and foods including onions, grapes, cauliflower, tomatoes, morning glory,
wild mustard, milk, beans, peas, cheese and chicken breast. Natural emissions from
a variety of tree species contain acetone vapour.

JD Reisman, Environmental Health Criteria for Acetone (Draft), Environmental Health
Criteria No. 207, International Programme on Chemical Safety (INCHEM) (1998) (see

WO 03/039483

PCT/US02/36028

Section 1 .4, "Environmental Levels and Human Exposure**) at

http://vvrww.inchem.org/documents/ehc/ehc/ehc207.hto. For example, acetone may form
chemically within an environment by atmospheric oxidation of plant terpenes (Fruekilde et

Living organisms may internally generate acetone via a number of enzymatic routes. In

performed by Mycobacterium spp., including M vaccae; by desulfonation of 2-
propanesulfonate, as may be performed by Rhodococcus spp. and Comamonas spp.,
10 including C. acidovorans\ and by decarboxylation of acetoacetate, as may be performed by
Clostridium spp., including C. acetobutylicum, C. butyricum, and C.
saccharoperbutylacetonium.

In the vertebrate animals, including humans, the most common route of acetone synthesis is
15 by ketone body formation. Ketone bodies are compounds produced from the oxidation of
lipids by the liver and used as an energy source when glucose is not readily available. The
main compounds classified as ketone bodies include acetoacetic acid, P-hydroxybutyric
acid, and acetone. Ketones are always present in the body, and their levels increase during
fasting and prolonged exercise. Oxidation of fatty acids in liver mitochondria produces
20 acetyl-coenzyme A, which can be further oxidized via the citric acid cycle or undergo a
process called ketogenesis. Ketogenesis occurs primarily when glucose is not available as
an energy source and converts acetyl-coenzyme A to acetoacetate or P-hydroxybutyrate.
The liver releases acetoacetate and P-hydroxybutyrate to the bloodstream where it is carried
to peripheral tissues and is used as an alternative energy source. Acetoacetate is a P-

25 ketoacid and slowly undergoes spontaneous non-enzymatic decarboxylation to acetone and .
C0 2 (Scheme 1).

al. (1998)).

microbes, acetone may be synthesized, for example: by oxidation of isopropanol, as may be

Scheme 1

C0 2

O
li

H3C C CH3

acetoacetate

acetone

WO 03/039483 PCT/US02/36028
In tetrapod vertebrates, including mammals, the acetone thus formed is detectable in
respiration as a result of blood gas exchange in the lung.

Breath acetone levels have been correlated with blood acetone levels and so may be used as
5 an accurate indicator of blood acetone content. Thus, elevated breath acetone levels have
been demonstrated, in clinical studies of otherwise healthy human subjects, to be a reliable
indicator of fat metabolism and projected weight loss. Acetone is present in human breath
at endogenous levels of about 0.2-0.5 ppm (v/v) and increases to and above 5-25 ppm for
otherwise healthy individuals on long term, low carbohydrate diets. Similarly, breath

10 acetone concentrations increase in individuals on a high-fat diet. Each of these dietary
conditions is called a "benign dietary ketosis." Breath acetone concentration likewise
increases during short-term fasting, a condition known as "fasting ketosis," and after
prolonged exercise, a condition known as "post-exercise ketosis." Under starvation
conditions (or long-term fasting) or in diabetics whose insulin levels drop too low, the

15 concentration of breath acetone may become abnormally high (up to 70 ppm or higher),
indicating conditions called, respectively, "metabolic ketoacidosis" or "diabetic
ketoacidosis," diabetic ketoacidosis being a potentially fatal condition. In each of these
conditions, elevated acetone levels can be detected in the breath of juveniles or adults.

20 In addition to acetone elevation by benign dietary, fasting, and post-exercise ketosis, and by
diabetic and metabolic ketoacidosis, other conditions and diseases can also generate
elevated blood acetone, and thereby elevated breath acetone, levels. For example, blood
acetone elevation is also observed in, for example: 1) the female reproductive cycle (for
example, during pregnancy or during the post-partum interval preceding resumption of

25 ovulation); 2) the neonatal stage of development; 3) hypoglycemia (for example,

hypoglycemia of childhood or hypoglycemia caused by eating disorders or prolonged
vomiting); 4) inborn metabolic diseases (for example, maple syrup urine disease); 5) liver
dysfunction (for example, end-stage liver disease or hepatic ischemia); 6) glucocorticoid
deficiency; 7) growth hormone deficiency; 8) acute pancreatitis resulting from viral

30 infection (for example, systemic cytomegalovirus infection); 9) treatment with nucleoside
analogs (for example, in anti-retroviral therapy for HIV); 10) isopropanol ingestion or
intoxication; 11) ethanol intoxication; and 12) salicylate intoxication. In these conditions,
too, acetone can be detected by, for example, breath analysis of children or adults.

WO 03/039483 PCT/US02/36028

Thus, detection of acetone can be useful in a number of medically important applications.
For example, medical reports have identified obesity as a primary risk factor in diabetes,
hypertension, coronary heart disease, hypercholesterolemia and stroke. In many cases of
5 obesity, a controlled weight loss program can reverse these serious life-threatening diseases.
Acetone is a metabolite that can be detected to monitor the progress in and compliance with
such a weight loss program. Similarly, detection of acetone can be used to alert diabetic
subjects to the onset of ketoacidosis or to obtain a preliminary indication of the need to
. diagnose a subject for any of the other medical conditions or diseases in which elevated
10 acetone may be found. Therefore, acetone is a key diagnostic metabolite that can be used as
a means to monitor diet compliance, weight loss progress, medical treatment regimen
compliance, diabetes, and health wellness in subjects of all ages.

B. The Field of Acetone Detection

15 Acetone can be detected in any of the above-described situations, by use of various means.
A wide variety of means for acetone detection are known in the art, including those for
detecting acetone from liquid solution (for example, blood and plasma analysis, urine
testing) and those for detecting acetone from gas mixtures (for example, breath sampling,
ambient gas monitoring). A broad assortment of different methodologies have been

20 employed in acetone detection, monitoring, and analysis. These methodologies include
those relying on, for example: color indicator, optical reflection, heat-of-combustion,
electrical resistance, gas chromatography (GC), liquid chromatography (LC), photometry,
colorimetry, ultraviolet spectrometry (UV), infra-red spectroscopy and spectrometry (IR),
microwave spectroscopy, and mass spectrometry (MS) technologies.

These technologies, and thus the methodologies employing them, vary in their specificity:
some detect a broad range of volatile organic compounds (VOCs); some detect either
ketones (and ketoacids) alone or both ketones (and ketoacids) and aldehydes; and some
detect acetone specifically.

In a first group of acetone detection methodologies, acetone is monitored by use of any of a
variety of technologies that detect a broad range of VOCs, examples of which technologies
include the following. Ambient gas monitors that detect a broad range of VOCs include, for

WO 03/039483 . PCT/US02/36028

example: the Drager Polytron SE Ex detector, which employs a catalytic, heat-of-
combustion "pellistor" type sensor (catalog no. 68 09 760; 0.60 kilograms); and the Drager
Polytron IR Ex gas detector, which uses an infrared sensor (catalog no. 83 12 550; 1 .9
kilograms) (both available from Drager Sicherheitstechnik GmbH, Liibeck, Germany).

Fluid-solid interaction-based detection of VOCs relies upon gas or liquid adsorption onto a
solid phase (optionally including a chemical derivatization reaction), followed by
colorimetric or photometric detection of the adsorbed and/or derivatized compound(s).
Such fluid-solid interaction technologies are discussed in British Patent No. 1 082525, which

1 0 discloses detection of organic compounds containing active or activated hydrogen atoms,
wherein metal zeolites are used as the solid. VOC detection involving liquid adsorption
onto solid and utilizing photometric detection is discussed in US Patent No. 4,882,499.
This patent teaches a liquid detector utilizing fiber optics to detect changes in optical
coefficient of reflection of liquid sample absorbed by capillary action; a hydrophobic

15 fibrous or sintered matrix is used as the adsorptive solid for this purpose.

In another approach to VOC detection, gas that adsorbs onto a solid is sensed by electrical
resistance/conductivity detection: this is described in US Patent No. 5,382,341. This patent
describes a process of manufacturing smoke-detecting elements in which a bismuth oxide
20 film is deposited onto a substrate layer, and, thereafter, is electrically connected to a means
for measuring resistance. In this case, the solid may comprise bismuth oxides, Bi-Fe-
oxides, Bi-V-oxides, or Bi-Mo-oxides. Similarly, German Patent No. 028062 describes a
gas adsorption method in which the solid comprises an adsorbent layer of a semiconductor
device.

A further methodology for VOC detection relies upon a gas phase chemical derivatization
(halogenation) reaction to produce a detectable halogenated product. Examples of such gas
phase derivatization methodologies for VOC detection are taught in US Patent No.
4,198,208 (describing reaction with chlorine and detection of chlorinated species) and in
30 German Publication No. 4007375. All of the technologies in this first group are capable of,
and taught for, detecting acetone, though non-specifically as a member of the class of
VOCs.

WO 03/039483 PCT7US02/36028
A second group of methodologies used for acetone detection are those that detect
. ketones/ketoacids or both ketones/ketoacids and aldehydes generally. The technologies
utilized in these methods depend on chemical derivatization of, for example, acetone, to
produce a colored product. The colored product can then be visually inspected to obtain a
5 qualitative result. Similarly, the colored product can be visually compared with a color
standard chart to obtain a semi-quantitative reading. Alternatively, the degree of coloration
can be quantitatively assessed by means of colorimetry or photometry. The most common
derivatization reactions employed in these technologies are those based on: 1) reaction with
salicylaldehyde; 2) reaction with hydrazine (or a phenylhydrazine, for example, 2,4-
10 dinitrophenylhydrazine); and 3) reaction with nitroprusside. Many other chemical
derivatization reactions are known, but are not commonly employed, for detection of
aldehydes and ketones/ketoacids (including acetone), because those reactions use high
temperatures or caustic, non-durable, or expensive reagents that make widespread use
impractical,

In salicylaldehyde-based assays, the ketone or ketoacid is introduced to an alkaline solution
containing salicylaldehyde, whereupon an orange or red derivative is produced. For
example, both acetone and acetoacetate may be detected in urine by this route, as disclosed
in U.S. Patent No. 2,283,262.

In hydrazine-based and phenylhydrazine-based assays, aldehydes, ketones, and ketoacids
are derivatized to form one or more hydrazone or phenylhydrazone compounds. For
example, gas phase acetone absorption into liquid solution for chemical derivatization by
this route is described: in US Patent No. 4,93 1 ,404, which discloses derivatization by
25 reaction with a hydrazine- or phenylhydrazine-coupled cation exchange matrix, followed by
colorimetric detection of the, for example, yellow, derivative; and in British Patent No.
2253910, which discloses derivatization by reaction with a hydrazine solution, followed by
electrical resistance detection of the derivative.

30 In nitroprusside-based assays, aldehydes, ketones, and ketoacids are reacted with

nitroprusside, that is a salt of nitroprussic acid (for example, sodium nitroprusside, that is
sodium nitroferricyanide), to form a derivative(s) that, in the presence of an amine, forms a
pink or purple complex. In some methods, the amine is present during the nitroprusside

WO 03/039483 PCT/US02/36028

reaction for immediate coloration, while in others, an amine-containing solution is added
later to develop the color. The nitroprusside reaction is the one most commonly used to
detect acetone in the context of personal health monitoring, for example, in diabetes or in
weight loss. Such nitroprusside methodology is typically found in one of three different
5 formats: gas sampling tubes, fluid testing strips, and fluid testing tablets.

A variety of nitroprusside tube assay devices are commonly used. One of the most common
is the Draeger tube (that is the DrSger tube), for example, the Draeger acetone detector tube
(catalog no. DRAG CH22901 from SAFECO, Inc., Knoxville, Tennessee). The Draeger

10 acetone detector tube can be used for breath analysis, but is mainly employed for ambient
gas sampling in which a pump draws an air sample through the tube. A similar assay
employs the Draeger Chip Measurement System (catalog no. 540-CMS from Safety First of
Middleton, WI; 0.74 kilograms) which utilizes a "chip," that is a planar, parallel array of
Draeger tubes of capillary dimension. This hand-held device pumps a gas sample into the

15 capillary tube(s), and the optics and electronics within the device perform colorimetry to
convert the degree of coloration of the derivative within the tube into a quantitative digital
signal. Similarly, methods for quantitatively monitoring acetone (and other ketones) in gas
samples by photometric detection of the colored derivative, are taught in manufacturer
information available with MSA acetone detector tubes (catalog no. 226620, available from

20 Ben Meadows Co., a subsidiary of Lab Safety Supply Inc., PO Box 5277, Janesyille, WI
53547). Also, US Patent No. 5,174,959 discloses a nitroprusside tube assay device
containing two solid matrices: a nitroprusside-coupled matrix and an amine-coupled
matrix. The device may be used with gas samples, in which case a solvent such as methanol
is added, or with liquid samples such as urine.

Nitroprusside fluid testing strips and tablets are typically marketed as urine ketone test strips
and tablets. Examples of such ketone test strip products are CHEMSTRIP K (produced by
Roche Diagnostics Corp., Indianapolis, Indiana) and KETOSTIX (produced by Bayer
Corp., Diagnostics Division, Tarrytown, New York). Exemplary ketone test tablets include
30 the AMES ACETEST reagent tablets (catalog no. AM-238 1 , available from Analytical
Scientific, Ltd., San Antonio, Texas).

WO 03/039483 PCT/US02/36028
A third group of methodologies are acetone-specific. These acetone-specific detection
methodologies utilize analytical devices and techniques. For example, acetone-specific
detection methodologies include: gas chromatography detection, described in A Amirav et
al., "Human Breath Analyzer for Medical Diagnostics," at
5 http://tau.ac.iiychemistry/amirav/breath.shtml (Nov 2000); liquid chromatography detection
using a micro-column, as disclosed in US Patent No. 6,063,283; and mass spectrometry
detection, as described in US Patent No. 5,999,886, for use in the context of acetone vapor
detection in semiconductor wafer processing chambers.

10 Specific detection of acetone in fluids by IR spectroscopy is disclosed in US Patent No.
5,355,425 (for liquids) and US Patent No. 4,587,427 (for gases). Specific detection of
acetone in gases by FTIR spectrometry is disclosed in RT Kroutil et al., "Automated
Detection of Acetone, Methyl Ethyl Ketone, and Sulfur Hexafluoride by Direct Analysis by
Fourier-Transform Infrared Interferograms," Applied Spectroscopy 48(6):724-32 (June

15 1994). Specific detection of acetone in gases by microwave spectroscopy is described in the
abstract of Medical Technology Co-Operation Offer 131.C, entitled "Microwave Gas
Spectroscopy for the Analysis of Exhaled Air," from the Nizhny Novgorod Region
Cooperation of the East- West Agency (OWA) of the Association for Innovation Research
and Consultation mbH of the Innovation Consulting Institute (InnovationsBeratungsInstitut,

20 Dusseldorf, DE) (See "Medical Technology" link at http://www.owa-ibi.com/owa-
deutsch/index.html).

In addition, a number of acetone-specific detection methods employing hyphenated
analytical techniques are also well known in the art. For example, a selection of such

25 methods, including those relying on GC-HPLC, GC-FID ("Flame Ionization Detector"),
GC-MS, GC-RGD ("Reduction Gas Detector"), and HPLC-UV techniques, are described in
JD Reisman, Environmental Health Criteria for Acetone (Draft), Environmental Health
Criteria No. 207, International Programme on Chemical Safety (INCHEM) ( 1 998) (see
Section 2, "Identity, Physical and Chemical Properties, and Analytical Methods") at

30 http://www.inchem.org/documents/ehc/ehc/ehc207.htm.

Thus, gas (into liquid) absorption with chemical reactions, gas (onto solid) adsorption with
and without chemical reactions, liquid (onto solid) adsorption with and without chemical

WO 03/039483 PCT/US02/36028
reactions, gas phase chemical reactions, solution phase chemical reactions, UV, IR, GC, LC,
MS, and other technologies have all been used for acetone detection. However, all of these
technologies have drawbacks.

5 For example, it is desirable, for environmental, health, and safety reasons, to select an
acetone detection method that is specific for acetone. This is especially important in the
area of subject self-monitoring, for example, for diabetes and for dieting. Thus, broad VOC
detection technologies are less desirable for these purposes. While the acetone-specific
methodologies currently in use are specific for acetone, they require bulky equipment that is

10 not easily transportable and is relatively expensive to obtain and maintain, and they are
impractical for use by individuals lacking medical or scientific training. It is desirable for
acetone detection technology to be light-weight, readily transportable, low cost, and easy-
to-use. These features are also especially important in the area of subject self-monitoring.
Thus, the currently available acetone-specific detection technologies are less desirable for

1 5 these purposes and reasons.

In contrast to these acetone-specific technologies, most of the currently available
methodologies that detect ketones/ketoacids or both ketones/ketoacids and aldehydes
generally are relatively inexpensive, light-weight, readily transportable, and easy-to-use.

20 Yet, without the use of a secondary detection system, such as colorimetry or photometry,
these tests produce only a qualitative or semi-quantitative result: a visualized color reading.
Second, these tests are not specific for acetone: the presence of acetoacetate, other ketones,
and aldehydes can result in a falsely intensified color reading. Finally, these tests are
susceptible of producing false positive and false negative results; the former falsely indicate

25 the presence of elevated acetone, the latter falsely indicate the absence of elevated acetone.
For example, in the most commonly used assays (nitroprusside assays): false positive
results often occur with subjects taking, for example, sulfhydryl drugs such as captopril, or
when other ketones or aldehydes are present; and false negative results often occur when
testing highly acidic samples, for example, samples of urine from subjects taking large

30 doses of vitamin C (ascorbic acid). It is desirable for acetone detection technology to be
reliable, to be specific for acetone, and to be capable of producing a directly quantitative
result. These features are also especially important in the area of subject self-monitoring.

WO 03/039483 PCT/US02/36028
Thus, the currently available ketone/ketoacid and aldehyde detection technologies are less
desirable for these purposes.

Therefore, a need exists in the field of acetone detection for an acetone detection technology
5 that is acetone-specific, light-weight, readily transportable, low cost, easy-to-use, reliable,
and capable of producing a directly quantitative result. It would also be advantageous for
such a technology to be capable of producing an electronic result, for example, a digital
result (or in the case of a photonic-type computer or other instrument, capable of producing
a photonic result). Affordable, disposable, specific, single-use devices for monitoring
1 0 acetone levels in biological samples by, for example, a subject at home, are not readily
available.

Other fields, such as the field of ethanol detection, utilize enzyme-based technologies.
Enzyme-based technologies can be analyte-specific and can take the form of light-weight,

15 readily transportable, low cost, easy to use, and quantitative devices. For example, in the
field of ethanol detection in biological samples, gas phase ethanol detection has been
performed by means of an enzyme-linked electrochemical sensor, using either alcohol
oxidase (AOX) or primary alcohol dehydrogenase (ADH) as the enzyme. In one,
exemplary ethanol detection system, a thick-film, screen-printed enzyme electrode using

20 ADH/NAD* immobilized in hydroxyethylcellulose is utilized for monitoring ethanol vapor.
This ethanol-detecting enzyme electrode is activated by dipping it into buffer, the ethanol-
containing sample is applied, and the resulting NADH produced by enzymatic action is
monitored amperometrically at 650 mV (vs. Ag/AgCl). A similar electrode has been
described for measuring ethanol vapor, wherein ADH is immobilized in reverse micelle

25 media. The ethanol is partitioned into the aqueous phase where it is effectively

concentrated in the medium where ADH can act upon it. These technologies are readily
adaptable to easily transportable, low-cost detectors that can be used to monitor or self-
monitor breath ethanol at home, at work, and in other environments remote from clinics and
testing laboratories. However, these devices are not designed for, and the enzymes utilized

30 (for example, primary alcohol dehydrogenase) are not capable of, acetone detection. Thus,
if it could be devised, an enzyme-based technology might be able to offer many of the
benefits needed in the field of acetone detection.

WO 03/039483 PCT/US02/36028
Nevertheless, all of the above-described acetone detection methods and devices rely on non-
enzymatic technologies. Enzymatic measurement of acetone in environmental and
biological samples has not been described to date. Thus, it appears that, in addition to the
need for an acetone detection technology having the advantageous features described above,
the field of acetone detection is also lacking the use of, and thus the benefits of, enzyme-
based technologies.

WO 03/039483

SUMMARY OF THE INVENTION

PCTAJS02/36028

The present invention is directed to enzyme-based detection systems that are acetone-
specific, including acetone-specific, enzyme-based detection systems that are light-weight,
5 readily transportable, low cost, easy-to-use, reliable, capable of producing a directly
quantitative result, and capable of producing an electronic result. The capability of
producing an electronic, for example, digital, result makes such a device readily adaptable
to computerized data collection and transmission via a communication network such as the
Internet (through either hard-wired or wireless means), including via the methodologies
10 described in WO 01/63277. Such computerized data collection and transmission would
permit methods of using the acetone-specific detectors of the present invention to be
especially useful for compliance monitoring, coaching, and instructing.

In comparison with the currently available, lightweight, readily transportable, low cost,
15 easy-to-use acetone detection technologies, the acetone-specific, enzyme-based detection
systems of the present invention also offer improved sensitivity of acetone detection. None
of the above-referenced documents suggests improving the sensitivity of acetone detection
via enzymatic methodologies. Enzyme-based measurement of acetone in mammalian
biological samples has not been described to date. However, as discussed above, specific
20 detection of acetone can be critically important in a wide variety of medical conditions, and
would be especially useful in the context of subject self-monitoring. However, such
acetone-specific detection technologies are not generally available outside the laboratory or
clinic setting, nor practical for regular subject self-monitoring. For these reasons, a need
exists to provide improved, less cumbersome, and easier-to-use methods and devices for the
25 specific detection of acetone in biological samples. Thus, detection of acetone by means of
acetone-specific enzyme-based detectors according to the present invention would be
particularly useful in a number of medically important applications.

Acetone-specific enzyme systems useful in quantitatively measuring acetone levels in
30 biological samples have been created to address the problems with the state of the art

technology. The enzyme systems and detectors of the present invention selectively act upon
acetone as a substrate. The term "selectively" is intended to characterize an enzyme,
system, or device that exhibits substantially greater affinity for or activity toward acetone

WO 03/039483 PCT/US02/36028
over other potentially available substrates, that is: an enzyme that is preferential for acetone
among other chemical species normally present in a sample, for example, human breath,
and/or an enzyme that (in the context of the acetone detectors of the present invention)
would not appreciably come into contact with other substrates significantly competitive
with acetone. Accordingly, in the systems and devices of the present invention, enzyme
interactions with substrates other than acetone or reactive acetone derivatives are negligible.

One or more enzyme systems may be combined in a device, such as a biosensor, to facilitate
sample measurement. Enzyme interaction with acetone may be directly or indirectly
detected by various detection methods known in the art. These enzyme systems are useful
in, for example, an in-home device for determination of acetone levels in human biological
samples, such as breath, saliva, or urine, thereby providing a non-invasive means for
monitoring subject wellness and/or for monitoring subject compliance with weight loss
diets, health management programs, and treatment regimens.

Accordingly, an aspect of the invention is an acetone-specific enzyme system that couples
enzyme-mediated metabolism of acetone to electrochemically detectable signals produced
via one or more of the signal mediators described hereinabove. Specifically, an acetone-
specific enzyme system, including an enzyme that selective targets acetone as a substrate,
coupled to a detectable signal mediator is a preferred aspect of this invention, especially
where the detection is performed by use of electrochemical or photometric means. Any
acetone-specific enzyme capable of linking to an electrochemically detectable co-factor or
by-product may be suitable for the enzyme system of this invention.

The acetone-specific enzyme system may be present in a lyophilized form until contacting
the biological sample. Moreover, the electrochemical biosensor, according to the invention
as described above, may comprise an acetone-specific enzyme system that is storage
stabilized by the presence of a disaccharide, such as trehalose. The biological sample may
be either in liquid or vapor form, but is preferably in vapor form.

An important feature for enzyme-mediated quantitation of acetone in samples such as
human breath is use of an enzyme system having high acetone sensitivity (for example, a
lower detection limit in the range of about 0.2-0.5 ppm (v/v)) that may be incorporated into

WO 03/039483 PCT/US02/36028
a device. Additionally the enzyme system must be robust, stable, and relatively
inexpensive. Optimally, the device should be capable of computer-interfacing, highly
selective, portable, and have low interference from other metabolites found in breath, such
as ethanol. Enzyme electrode technology (biosensors) is particularly suitable to meet these
5 requirements, where selective enzymatic reactions are involved, due to the sensitivity of
detection and the relatively simple instrumentation needed. Electrochemical detection
depends on the direct measurement of current generated by the reaction of the detected
species at the electrode. Coupling oxidoreductase (redox) enzyme reactions to electrodes
has been an attractive approach to developing biosensors. In particular, electrochemical
10 detection of reduced nicotinamide adenine dinucleotide (NADH) or hydrogen peroxide has
been utilized in amperometric biosensors for a range of substrates, namely glucose {that is
the reactions of glucose dehydrogenase and glucose oxidase respectively).

The acetone-specific enzyme system preferably contains an acetone-specific enzyme
15 selected from the group consisting of acetone mono-oxygenase, acetone carboxylase, and
secondary alcohol dehydrogenase. In particular, the inventive acetone-specific enzyme
system preferably contains an acetone-specific enzyme system selected from the group
consisting of acetone carboxylase product formation coupled to NAD(P)H oxidation,
acetone carboxylase ATP-hydrolysis coupled to NAD(P)H oxidation, acetone carboxylase
20 ATP hydrolysis coupled to H2O2 formation, secondary alcohol dehydrogenase (S-ADH)
coupled to NAD(P)H oxidation, S-ADH catalyzed NAD(P) + formation coupled to H 2 0 2
production, and acetone mono-oxygenase coupled to NAD(P)H oxidation.

In still another preferred embodiment, the acetone-specific enzyme system according to the
25 invention contains a signal mediator selected from the group consisting of organic cofactors,
inorganic cofactors, multi-electron transfer mediators and enzyme reaction by-products.
Preferably, a signal from the electrochemically detectable signal mediator is linearly or
exponentially amplified by magnifying electrochemical signal output via recycling enzyme
substrates.

In a preferred embodiment of the invention, the acetone-specific enzyme system contains an
acetone-utilizing enzyme obtained from a vertebrate or microbe, more preferably from a
mammal, fungus, bacterium, or Archaeon.

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PCTYUS02/36028

In another preferred embodiment of the invention, the acetone-specific enzyme system
contains secondary alcohol dehydrogenase obtained from a mammal or from a
microorganism selected from aerobic and anaerobic bacteria, yeast, fungi, and
methanogenic Archaea. Preferably, a secondary alcohol dehydrogenase is isolated from a
species of Thermoanerobium, Xanthobacter, Pseudomonas, Rhodococcus, or
Mycobacterium; especially preferred is a secondary alcohol dehydrogenase isolated from
Xanthobacter autotrophicus strain Py2.

In yet another preferred embodiment of the invention, the acetone-specific enzyme system
contains acetone carboxylase isolated from a species of aerobic or anaerobic bacteria, more
preferably from a species of Xanthobacter, Rhodococcus, or Rhodobacter.

In a preferred embodiment of the invention, an electrochemical biosensor for detecting
acetone in a biological sample contains at least one acetone-specific enzyme system as
generally described above, and a means for detecting a product resulting from a reaction
between the at least one acetone-specific enzyme system and acetone in the biological
sample. The acetone-specific enzyme system of the biosensor preferably comprises an
enzyme selected from the group consisting of acetone carboxylase, secondary alcohol
dehydrogenase, and acetone mono-oxygenase. In particular, the acetone-specific enzyme
system may comprise at least one member selected from the group consisting of: 1)
secondary alcohol dehydrogenase (S-ADH)-catalyzed reduction of acetone, with
concomitant NAD(P)H consumption (oxidation); 2) acetone carboxylase reaction coupled to
P-hydroxybutyrate dehydrogenase consumption of NAD(P)H; 3) acetone carboxylase
reaction ATP hydrolysis coupled to NAD(P)H consumption; 4) S-ADH reaction NAD(P) +
formation coupled to H 2 0 2 formation; 5) acetone carboxylase reaction ATP hydrolysis
coupled to H 2 0 2 formation; 6) acetone carboxylase reaction coupled to P-hydroxybutyrate
dehydrogenase NAD(P) + formation coupled to H 2 0 2 formation; 7) acetone mono-oxygenase
coupled to NAD(P)H oxidation; 8) acetone mono-oxygenase coupled to H 2 0 2 formation;
and 9) acetone monooxygenase-catalyzed NAD(P)+ formation coupled to H202 formation.

In manufacturing a biosensor containing one or more acetone-specific enzyme systems,
various enzymatic by-products and/or factors may be employed for the production of

WO 03/039483 PCT/US02/36028

electrochemical signals for detecting enzymatic reactions. Organic cofactors, such as NAD,
NADH, NADP, NADPH, FAD, FADH, FMN, FMNH, Coenzyme A, Coenzyme Q, TTQ
(Tryptophan Tryptophylquinone) and PQQ (Pyrroloquinolinequinone), may be used.
Electron transfer mediators may be employed to improve the kinetics of electron transfer,
5 since organic cofactors sometimes easily foul a detector. Mediators useful in multi-electron
transfers for reduced forms of cofactors may be included in the enzyme systems of the
present invention. Similarly, inorganic cofactors or indicators may be employed for the
production of electrochemical signals. Enzymatic reaction by-products, such as hydrogen
peroxide or ammonium, and energetic molecules may also be used in the invention for
10 coupling acetone metabolism to electrochemically measurable signals. Combinations of
these cofactors and systems may be used.

In a preferred embodiment, the electrochemical biosensor according to the invention may
further comprise a means of interfacing with an analytical device, such as a computer linked
15 to the Internet.

In a preferred embodiment, the electrochemical biosensor according to the invention may be
characterized in that two or more acetone-specific enzyme systems are present. In such a
biosensor, the two or more acetone-specific enzyme systems may be disposed on separate

20 electrodes, for example, electrodes grouped in sequentially arranged clusters along a sample
detection pathway, forming an electrode array. In other words, electrodes intended for
contact with a single acetone-containing sample may be grouped within a common sample
pathway. Several groups having the same or different acetone-specific enzyme systems
may be arranged sequentially along the pathway to enhance the sensitivity of acetone

25 detection.

In an alternative preferred embodiment of the electrochemical biosensor of the invention,
one or more of such separate electrodes in an array may be acetone-specific, while one or
more additional separate electrodes in the array may be capable of detecting a non-acetone
30 analyte. For example, in an ethanol detector in which acetone, if present, may be

secondarily detected by an electrode designed for ethanol detection, the inclusion of a
separate, acetone-specific electrode can provide a basis for correcting for such acetone -
interference. In another example, detection of both acetone and at least one further analyte

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may be desired in order to determine a useful ratio of analyte concentrations, which ratio
may be desirable for detecting, for example, a medical condition or disease state, such as
diabetes: in diabetes and other conditions, the ratio of acetone to, for example, 0-
hydroxybutyrate, may be useful to provide a more complete indication of the subject's
5 metabolic state. The device of which the electrode array is a part, or an instrument to which
the device can be interfaced, may electronically generate separate results for each electrode
or separate results for each analyte. Alternatively, some type of combined result may be
generated, for example, a sum, a difference, a ratio, etc. Electronics within the device, or
electronics within a computer or other electronic instrument to which the electrode may be

10 interfaced, can utilize the result(s) from the acetone electrode(s), for example: to calculate a
ratio between the acetone and the other measured analyte(s) (for example, 0-
hydroxybutyrate); or to calculate a subtractive correction factor for interference of acetone
in the results(s) obtained from the other array electrode(s) designed for detection of non-
acetone analyte(s) (for example, ethanol) and then calculate a corrected reading therefor.

15 One embodiment of such an array-based electronic (or electronically-coupled) device is an
"electronic nose," which may be used for, for example: diagnosis of medical condition
from breath analysis; diagnosis of medical conditions from gases or vapors emitted by
infected fluid samples, for example, bacterially infected urine; detection of tank car and
other industrial container gas leakage; quality control monitoring of fermentations and food
20 processing systems by detecting emitted vapors; and monitoring of enclosed air space
quality.

In a preferred embodiment of the invention, a method of detecting acetone in a biological
sample involves introducing a biological sample containing acetone to a biosensor
containing at least one acetone-specific enzyme system that utilizes acetone as a substrate,
and detecting the interaction between the acetone and the acetone-specific enzyme system.
The biological sample is preferably a vapor sample. Detection may be achieved via
photometric, calorimetric, or electrochemical means. The method may further comprise
facilitating electrochemical transduction of the at least one acetone-specific enzyme system
and the acetone in the vapor sample via an electrochemical ly treated electrode, and
electrochemically detecting a product resulting from a reaction of the at least one acetone-
specific enzyme system with the acetone in the vapor sample.

WO 03/039483 PCT/US02/36028
The at least one enzyme system used in this method may comprise a member selected from
the group consisting of acetone mono-oxygenase, acetone carboxylase and secondary
alcohol dehydrogenase. Specifically, the enzyme system may include any one or more of:
1) secondary alcohol dehydrogenase (S-ADH)-catalyzed reduction of acetone, with
5 concomitant NAD(P)H consumption (oxidation); 2) acetone carboxylase reaction product
formation coupled to (for example, p-hydroxybutyrate dehydrogenase) consumption of
NAD(P)H; 3) acetone carboxylase reaction ATP hydrolysis coupled to NAD(P)H
consumption; 4) S-ADH reaction NAD(P) + formation coupled to H2O2 formation; 5)
acetone carboxylase reaction ATP hydrolysis coupled to H2O2 formation; 6) acetone

10 carboxylase reaction coupled to P-hydroxybutyrate dehydrogenase NAD(P) + formation
coupled to H 2 0 2 formation; 7) acetone mono-oxygenase coupled to NAD(P)H oxidation; ;
8) acetone mono-oxygenase coupled to H2O2 formation; and 9) acetone monooxygenase-
catalyzed NAD(P)+ formation coupled to H202 formation. The method preferably
comprises electrochemically detecting acetone in the vapor sample at a level of 0.5 ppm to

15 10 ppm.

Several areas of subject care may be enhanced by using acetone-specific enzyme systems
for monitoring acetone levels. In this regard, a home biosensor capable of enzyme-
mediated acetone detection in biological samples would permit subjects, care-givers, and

20 support groups to closely manage weight loss programs. Similarly, the inventive biosensor
would enable subjects suffering from acetone-related conditions, such as diabetes, to
monitor weight loss and/or the onset of ketoacidosis, as well as to non-invasively manage
their diet and medications, thereby controlling acetpne production. Another use for an
acetone-specific enzyme system in a sensor is remote prescription medicine management

25 through tagging of drug formulations with acetone or acetone-producing compounds.

In a preferred embodiment, an electrochemical biosensor device of the present invention
uses enzymes specific for acetone as a substrate, wherein an acetone-enzyme system
reaction is linked to a means of electrochemical detection. Use of such an acetone-specific
30 monitoring device requires that the biosensor be able to detect low levels of acetone,

particularly when the acetone to be monitored is contained in vapor samples. For example,
electrochemical acetone biosensors according to the invention may be used to assist subjects
in management of diabetes by measuring acetone levels in biological samples. Ketoacidosis

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readings and warnings may be obtained quickly by subjects at home, by incorporating into
the device either means for generating (from the electrochemical response present during
acetone detection) a qualitative or quantitative result, or means for interfacing the device
with a computer, instrument, or apparatus containing such means for generating. One
example of means for generating such a qualitative or quantitative result is electronic
circuitry. Electronic circuitry may be incorporated into an acetone-detecting biosensor
device according to the present invention, or by the biosensor may be designed to be
capable of interfacing with a computer or other electronic instrument containing such
circuitry. One such aspect uses a signal from the electrochemically detectable signal
mediator that is amplified by recycling enzyme substrates to multiply electrochemical signal
output. Such circuitry would be selected, for example, to be capable of generating and/or
graphically/textually/symbolically displaying a qualitative (for example, "low," "moderate,"
"high," "safe," or "danger") or a quantitative (for example, "10 percent (%) over target,"
"6% below average," "12 ppm," or "40 mg/dL") result or results. The device or instrument
may also contain a data storage means to record one or more test results. The device can
also be interfaced with a computer network, such as the Internet, in order to transmit
readings generate by the device to, for example, a clinic, laboratory, doctor's office, or
support group, for example, as disclosed in WO 01/63277. Furthermore, obese subjects
seeking to monitor weight loss may do so utilizing acetone-specific biosensors according to
the invention in the privacy of their own homes, or with the remote help of medical
professionals, caregivers, or support groups. This aspect of the invention is facilitated by
the fact that ketosis is considered the best indicator of successful dieting, and acetone levels
have been correlated to the component of pound weight loss due to fat.

Acetone-specific enzyme systems and sensors/electrodes of the present invention may be
used to monitor ketogenic diet-utilizing subjects for seizure control, to detect gestational
diabetes, to aid in Type I diabetes monitoring or Type II diabetes management, to monitor
client progress in weight loss and eating disorders counseling and in high performance
fitness training, and to assist in livestock management.

In addition to monitoring weight loss and managing acetone-related diseases, another
medical application of an acetone-specific enzyme system according the invention is
monitoring the availability of pharmaceutical compounds administered to a subject. The
release into the bloodstream of therapeutics tagged with acetone or an acetone-producing

WO 03/039483 PCT/US02/36028
compound could be tracked within a subject using a biosensor comprising an acetone-
specific enzyme system as described herein. For example, an orally active therapeutic could
be formulated with a tag comprising acetone, or acetoacetatic acid, or, for example, an
acetoacetate salt, ester, or amide. Acetoacetate spontaneously decarboxylates to release
5 acetone. Upon ingestion of the "tagged" therapeutic, the release of acetone from the

formulation into the subject's bloodstream could be monitored by a biosensor comprising an
acetone-specific enzyme system. Levels of acetone detected in the breath of a subject
would then be detected by photometric or electrochemical means via oxidoreductase
reactions linked to the acetone-specific enzyme system. Preferably, the dosed compound is
10 administered parenterally, for example, by injection or implant placement, but other, non-
parenteral routes of administration, for example, oral administration, are also suitable.

Thus, in another preferred embodiment of the invention, a method of using an acetone-
specific enzyme system involves combining acetoacetate or a pharmaceutically acceptable

15 salt, ester, amide, or other suitable derivative thereof, with a pharmaceutical compound,
thereby tagging the compound; administering the tagged compound to a subject; and
thereafter, monitoring release of the tagged compound into the subject's bloodstream via
detection of acetone in a subject's biological sample by means of the inventive acetone-
specific enzyme system. Specifically, release of the tagged compound is followed via

20 measuring acetone results from acetoacetate breakdown, which acetone is detected in a
biological sample by acetone interaction with the acetone-specific enzyme system of the
invention. Multiple readings by the present enzyme system can thereby be used to monitor
the rate of release of acetoacetate, and thus the rate of disintegration of the administered
composition, which correlates with the rate of release of the pharmaceutical active

25 ingredient(s) to the biological system (that is its bioavailability). Verification of drug

delivery and of subject compliance with a drug treatment regimen may be performed in this
way.

In another preferred embodiment, a method of measuring biodegradation of materials
30 involves tagging a biocompatible implant or device by combining acetoacetate, or a

pharmaceutically acceptable salt, ester or amide derivative thereof, with a biocompatible
composition to form a biocompatible implant or device; placing the biocompatible implant
or device in a biological system (for example, a cell culture, biotic environment, animal or

WO 03/039483 PCT/US02/36028
human subject); and measuring release of acetone produced by degradation of acetoacetate
released by erosion or degradation of the biocompatible implant or device. The
biocompatible implant or device may be simply a sample of a material being tested for
erosion or degradation in the biological system, or it may be an operative implant or device,
for example, containing a pharmaceutical active ingredient whose controlled release from
the implant or device is desired. Release of the acetoacetate can be detected upon its
degradation to acetone, which interacts with an acetone-specific enzyme system of the
present invention. The amount of acetone measured can be correlated with the amount of
acetoacetate released of the implant, which in turn is indicative of decomposition of the
biocompatible implant within the body. Multiple readings by the enzyme system of the
present invention can thereby be used to monitor the rate of release of acetoacetate, and thus
the rate of erosion or degradation of the material of which the implant or device is
composed.

Biocompatible materials are those materials that do not impair normal biological ftinctions.
Thus, materials that are biocompatible are not, for example, biochemical inhibitors or toxic,
mutagenic, or carcinogenic compounds, and do not cause, for example, immune response,
allergic reaction, or blood clotting activity. The class of biocompatible materials includes
both biodegradable and non-biodegradable materials. Biodegradable materials decompose
in contact with a biological organism, either externally (for example, in the case of
microbial biodegradation) or internally (for example, in the human body). Exemplary
biocompatible, biodegradable polymers include, for example, proteins (for example,
collagen), polysaccharides and derivatives (for example, chitosan), polydioxanone, and
polyhydroxalkanoates (PHAs, for example: PHA polymers such as polyglycolic acid,
polylactic acid, polyhydroxybutyric acid, polyhydroxyvaleric acid; and PHA copolymers).
Other exemplary biocompatible polymers include polyethyleneglycol, polycaprolactone,
cellulosic polymers, polyvinylalcohol, polyhydroxyethylmethacrylate, and
polyvinylpyrrolidone. Further biocompatible materials include hydroxyapatite, ceramics,
glasses, various metals, and composite materials. New biocompatible materials are being
continually developed, including those intended for short-term (for example, weeks), long-
term (for example, months or years), and very-long-term (for example, years or decades)
use in a biotic environment. New compositions intended for use as biocompatible materials
must be tested to determine whether they are in fact biocompatible and to assess the half-life

WO 03/039483 PCT7US0 2/36028

of the composition in order classify it, for example, as a short-term or long-term
biodegradable material, or as a non-biodegradable material. The enzyme system of the
present invention may be used to determine the half-life of such a material.

5 In addition, the acetone-specific enzyme systems and sensors/electrodes according to the
present invention may be used in conjunction with detection means for other biological
components in order to assist in identification of bacterial infection (for example, in
conjunction with detection means for nitrous oxide, acetoin, and/or other biological
compounds) or to assess oxidative stress or early cancer detection.

In a preferred embodiment of the invention, a kit for detecting acetone in a sample includes
an acetone-specific enzyme system and a housing for the acetone-specific enzymes system,
wherein the housing has a port for introducing a sample to the acetone-specific enzyme
system. The acetone-specific enzyme system may be disposed on a disposable strip that fits
15 into the housing, or the acetone-specific enzyme system may be fashioned into the housing
to form a single disposable unit within the kit.

The present invention also provides a secondary alcohol dehydrogenase enzyme that is a
protein obtained fromXanthobacter autotrophicus Py2 (ATCC PTA-4779), having NAD + -

20 dependent secondary alcohol dehydrogenase activity, having the ability to reduce acetone to
isopropanol, having specific activity for ketones and secondary alcohols; having, for the
oxidation of isopropanol to acetone, (1) a pH optimum of approximately 7.8, and having,
for the oxidation of alcohols, (2) an average specific activity ratio for secondary-to-primary
alcohols of at least 50:1 when tested at pH 7.8 under equivalent conditions individually with

25 C3-C5 straight chain secondary alcohols and with C2-C5 straight chain primary alcohols;
having, for the reduction of acetone to isopropanol, (3) a pH optimum of approximately 6.2,
(4) an apparent K m of approximately 1 44 ±18 (iM, (5) an apparent F max of approximately
43.4 ±1.2 nmol acetone reduced-min^mg" 1 protein, (6) an apparent k^t of approximately
30.4 sec* 1 , (7) an apparent k c JK m of approximately 2.1 x 10 s , and (8) a K m for NADH of

30 approximately 5.1 ± 0.4 |iM; and comprising at least one polypeptide molecule that has (a)
a molecular mass of approximately 37.1 kDa as determined by mass spectrometry, (b) a pi
of approximately 7.4 as determined by isofocusing electrophoresis, and (c) a tetradecameric
N-terminal amino acid sequence of SEQ ID NO:7, and that is capable of being degraded to

WO 03/039483 PCT/US02/36028

form fragments having the amino acid sequences of SEQ ID NO: 8 to SEQ ID NO: 19. Also,
provided is the polypeptide thereof.

5 BRIEF DESCRIPTION OF THE DRAWINGS

Figure 1 graphically depicts electrochemical detection of acetone-dependent
oxidation of NADH catalyzed by P-450 monooxygenase.

Figure 2 depicts a sodium-dodecyl-sulfate-Polyacrylamide Gel Electrophoresis
1 0 (SDS-PAGE) analysis of S-ADH purification from X autotrophic™ st. Py2.

Figure 3 shows substrate specificity of S-ADH from X autotrophics strain Py2 in
performing ketone reductions.

Figure 4 shows substrate specificity of S-ADH fromZ autotrophicus strain Py2 in
performing alcohol oxidations. (NA indicates no activity was detected).
1 5 Figure 5 graphically depicts the stability of lyophilized S-ADH from X

autotrophicus st. Py2 stored at room temperature, alone and with various additives (10% by
weight trehalose, citrate, or sucrose).

Figure 6 is a schematic depiction of an acetone electrochemical sensor using S-
ADH. "

20 Figure 7 graphically shows a correlation of electrochemical response (o) and UV

spectrophotometry response (•) data of S-ADH-catalyzed reaction.

Figure 8 graphically depicts electrochemical detection of acetone-dependent
oxidation of NADH catalyzed by S-ADH from X autotrophicus Py2.

Figure 9 schematically depicts electrochemical detection of acetone-dependent
25 oxidation of NADH catalyzed by S-ADH from X autotrophicus Py2 using a carbon
electrode modified with the mediator Meldola's Blue (MB).

Figure 10 graphically presents the results of an electrochemical assay of NADH
concentration using a glassy carbon electrode modified with Meldola's Blue.

Figure 1 1 graphically presents the results of an electrochemical assay of acetone-
30 dependent NADH consumption catalyzed by S-ADH using a Meldola's Blue modified
screen-printed carbon electrode MB-SPCE.

WO 03/039483 PCT/US02/36028
Figure 12 graphically presents the results of an electrochemical assay of acetone-
dependent NADH consumption catalyzed by S-ADH using a glucose commercial test strips
and monitor.

Figure 1 3 is a schematic of electrochemical detection of S-ADH reaction coupled to
5 H2O2 formation specifically showing S-ADH coupled to lactate dehydrogenase (LDH) and
pyruvate oxidase (PO).

Figure 14 graphically depicts acetone-dependent H2O2 formation using S-ADH
coupled enzyme system.

Figure 15 is a schematic illustrating a general scheme for coupling S-ADH reaction
10 to H2O2 formation.

Figure 16 graphically depicts the results of a reflectance photometry assay of
acetone-dependent H 2 0 2 formation using S-ADH coupled enzyme system using glucose test
strips and monitor.

Figure 17 graphically depicts the results of an electrochemical assay of acetone-
15 dependent H2O2 formation using S-ADH coupled enzyme system using a disk platinum
electrode.

Figure 18 graphically depicts the stoichiometric results of an electrochemical assay
of acetone-dependent H2O2 formation using S-ADH coupled enzyme system.

Figure 1 9 graphically depicts the results of an electrochemical assay of acetone-
20 dependent H2O2 formation using S-ADH coupled enzyme system using a disposable
platinized carbon electrode.

Figure 20 graphically depicts the results of an electrochemical assay of acetone-
dependent H 2 0 2 formation using S-ADH coupled enzyme system using a disposable
platinized carbon electrode.
25 Figure 2 1 graphically depicts the results of an electrochemical assay of acetone-

dependent H2O2 formation using S-ADH coupled enzyme system using a disposable carbon
electrode embedded with the mediator cobalt phtalolocyanine.

Figure 22 schematically depicts the test gas sampling system employed to analyze
acetone from synthetic breath.
30 Figure 23 graphically depicts electrochemical detection of acetone using an S-ADH

coupled enzyme system, after partitioning from the gas phase to wetted foam.

Figure 24 graphically depicts electrochemical detection of acetone using an S-ADH
coupled enzyme system, after partitioning from the gas phase to an aqueous thin film.

WO 03/039483 PCT/US02/36028
Figure 25 schematically depicts electrochemical detection of acetone carboxylase
activity coupled to P-hydroxybutyrate dehydrogenase activity.

Figure 26 graphically presents the results of a spectrophotometric assay of acetone-
dependent NADH consumption using AC/HBDH coupled enzyme system.
5 Figure 27 schematically illustrates electrochemical detection of acetone carboxylase

(AC) reaction coupled to pyruvate kinase (PK), myokinase (MK), and lactate
dehydrogenase (LDH) activities.

Figure 28 schematically shows electrochemical detection of acetone carboxylase
(AC) reaction coupled to pyruvate kinase (PK), myokinase (MK), and pyruvate oxidase
10 (PO) activities.

Figure 29 presents a plot of spectrophotometric absorbance detection of H 2 0 2
generated in response to acetone by a coupled enzyme system containing acetone
carboxylase from Xanthobacter Py2, myokinase, pyruvate kinase, and pyruvate oxidase, in
the presence of horseradish peroxidase (HRP) and electron acceptor dyes; the HRP and dyes
15 permitting the photometric detection of H2O2.

Figure 30 schematically shows acetone-dependent H2O2 formation using an acetone
carboxylase coupled enzyme system; the result is electrochemical detection of acetone using
the acetone carboxylase (AC) reaction coupled to P-hydroxybutyrate dehydrogenase
(HBDH), lactate dehydrogenase (LDH), and pyruvate oxidase (PO) activities.
20 Figure 3 1 schematically shows electrochemical detection of P450 monooxygenase

coupled to galactose oxidase activities.

Figure 32 schematically illustrates signal amplification strategy using acetone
carboxylase (AC) coupled enzyme system containing acetoacetate decarboxylase (ACD),
myokinase (MK), pyruvate kinase (PK), and pyruvate oxidase (PO).
25 Figure 33 schematically illustrates signal amplification strategy using an acetone

carboxylase (AC) coupled enzyme system containing myokinase (MK), pyruvate kinase
(PK), lactate dehydrogenase (LDH), and lactate oxidase (LO).

Figure 34 schematically illustrates linear signal amplification strategy using a
secondary alcohol dehydrogenase and a pyrroloquinolinequinone-dependent alcohol
30 dehydrogenase (PQQ).

Figure 35 schematically illustrates linear signal amplification strategy using S-ADH
and secondary alcohol oxidase (SAO).

WO 03/039483 PCT/US02/36028

Figure 36 schematically illustrates exponential signal amplification utilizing S-ADH
coupled to beta-hydroxbutyrate dehydrogenase (HBDH), acetoacetate decarboxylase
(ACD), and SAO

Figure 37 schematically illustrates a linear signal amplification strategy using two
different pyridine nucleotide-dependent secondary alcohol dehydrogenases: S-ADH- 1
(NADPH-specific secondary alcohol dehydrogenase), S-ADH-2 (NADH-specific secondary
alcohol dehydrogenase), along with D (NADH-specific diaphorase), and MED (an electron
mediator).

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PCT/US02/36028

DETAILED DESCRIPTION OF PREFERRED EMBODIMENTS

Definitions

5 NAD(P) + is used herein to mean "either or both of NAD + (nicotinamide adenine

dinucleotide, oxidized form) and NADP* (nicotinamide adenine dinucleotide phosphate,
oxidized form)." NAD(P)H is used herein to mean "either or both of NADH (nicotinamide
adenine dinucleotide, reduced form) and NADPH (nicotinamide adenine dinucleotide
phosphate, reduced form)." Nicotinamide adenine dinucleotide is also called 3-carbamoyl-
10 l-p-D-ribofuranosyl-pyridinium hydroxide 5'-ester with adenosine 5 '-pyrophosphate, inner
salt. Nicotinamide adenine dinucleotide phosphate is also called 3 -carbamoyl- 1-|}-D-
ribofuranosyl-pyridinium hydroxide 5'^5*-ester with adenosine 2'-(dihydrogenphosphate)
5 '-(trihydrogen pyrophosphate), inner salt.

As used herein, " Annn" indicates "absorbance measured at NNN nanometers wavelength."
15 As used herein, "ennn" indicates "extinction coefficient measured at NNN nanometers
wavelength."

"Enzyme" as used herein means "catalytically functional biomolecule;" thus any
biomolecule that can perform a named catalytic function as its primary catalytic activity is
considered an enzyme of that name, regardless of other considerations such as origin, native
20 or engineered structure, size, etc.

"Platinized carbon," as used herein, indicates platinum-coated carbon, for example at least
partially platinum-coated carbon nanoparticles.

"Photometric," as used herein, indicates any detection mode in which photons are utilized
and includes, but is not limited to, colorimetric, spectrometry, spectrophotometry,
25 luminescence-based, chemiluminescence-based, electrogenerated chemiluminescence-
based, bioluminescence-based, and fluorescence-based methods.

In order to address certain difficulties associated with subject health maintenance, an
enzyme-based biosensor has been developed, which enables the coupling of enzyme-
30 mediated metabolism of acetone to electrochemically detectable signals produced via one or
more of the signal mediators. Any acetone-specific enzyme capable of linkage to an
electrochemically detectable co-factor or by-product may be suitable for the enzyme system
of the invention. In a preferred embodiment, an electrochemical biosensor for detecting

WO 03/039483 PCTAJS02/36028
acetone in a biological sample contains at least one acetone-specific enzyme system, and a
means for detecting a product resulting from a reaction between the at least one acetone-
specific enzyme system and acetone in the biological sample. The detection means may be
either electrochemical or non-electrochemical.

Acetone-Specific Enzymes

A number of enzymes, mainly from bacterial sources, have been described which
specifically utilize acetone as a substrate. These enzymes have been obtained from and/or
characterized in aerobic and anaerobic bacteria that are able to grow using acetone as a sole
10 carbon and energy source.

Acetone may be formed in bacteria by the action of secondary alcohol dehydrogenase (S-
ADH), an enzyme that operates in conjunction with one of two different acetone metabolic
pathways: an (^-dependent (oxygen utilizing) pathway in which the acetone is then

15 oxidized to produce acetol, and a C02-dependent (carbon dioxide utilizing) pathway in
which the acetone is then converted to acetoacetate. The acetone formation reaction
catalyzed by S-ADH is freely reversible and normally requires a coenzyme that is typically
either NAD(H) or NADP(H). The reduction of acetone to isopropanol by oxidation of
NAD(P)H (the reverse, S-ADH-catalyzed reaction) involves redox chemistry by which

20 acetone concentration can be monitored (for example, by means of electrochemical

determination of NAD(P)H consumption). A variety of secondary alcohol dehydrogenases
have been purified and characterized. Those best studied are S-ADHs obtained from
hydrocarbon oxidizing (that is propane utilizing) bacteria, which employ (^-dependent
acetone metabolic pathways. S-ADH enzymes have also been isolated from or described in

25 microorganisms not associated with hydrocarbon oxidation (that is propane degradative

metabolism). These include methylotrophic bacteria and yeast, methanogenic Archaea, and
fermentative anaerobes. Of these enzymes, S-ADH from Thermoanaerobium brockii is
commercially available as a heat-treated crude preparation or in purified form (available
from Sigma Chemical Co., St. Louis, Missouri). This enzyme is well characterized and is

30 an NADPH-specific dehydrogenase.

In some propane-oxidizing bacteria, acetone is formed as an intermediate that is then
understood to undergo hydroxylation in an 02-dependent mono-oxygenase-catalyzed

WO 03/039483 PCT/US02/36028
reaction to form acetol (hydroxyacetone). Acetol is then further oxidized to methylglyoxal
catalyzed by an acetol dehydrogenase, or is involved in a carbon-carbon cleavage reaction
producing CI and C2 fragments. Acetone mono-oxygenase, which is a pyridine nucleotide-
dependent enzyme, provides the necessary requirements for electrochemical detection in an
5 acetone biosensor (as described above). Acetone metabolism via acetol as an intermediate
has been identified in in vivo studies of acetone-utilizing bacteria. Also, P450 mono-
oxygenases have been identified in mammals as using an identical mechanism (to oxidize
acetone to acetol). An acetone mono-oxygenase suitable for use in an acetone-specific
enzyme system is a cytochrome P450 acetone mono-oxygenase isolated from mice (Mus

10 musculus). This monooxygenase has been reported as utilizing acetone as a substrate to
produce acetol, and is commercially available from PanVera Corporation (Madison, WI).
See F. Y. Bondoc et aL, Acetone catabolism by cytochrome P450 2E1 : Studies with
CYP2E1 -null mice. Biochemical Pharmacology, 58:461-63 (1999). The enzyme
responsible for this activity in bacteria has not yet been fully characterized. In addition,

15 acetone mono-oxgenase can be coupled to H 2 0 2 generation by including a galactose oxidase
in the enzyme system; galactose oxidase oxidizes acetol to form H 2 0 2 which can be
detected either electrochemically or non-electrochemically.

Mammalian P450 cytochromes containing acetone mono-oxygenase activity and P450
20 reductase, may be prepared from heptatic microsomes. P450 acetone mono-oxygenase
catalyzes the following hydroxy lation reaction:

NAD(P)H + H + + acetone + 0 2 -» NAD(P) + + acetol + H 2 0 (P450 acetone mono-
oxygenase)

P450 monooxygenases are typically comprised of two enzyme components including a
pyridine nucleotide-dependent reductase and an active site-containing oxygenase
component. N AD(P)H provides the necessary reductant for 0 2 activation and incorporation
of one oxygen atom into the aliphatic hydrocarbon substrate. With some P-450
30 monooxygenases, a third electron transfer component, cytochrome b$, will stimulate
activity. Acetone-dependent consumption of NAD(P)H by an acetone mono-oxygenase
reaction could be monitored electrochemically as described below for secondary alcohol
dehydrogenase-coupled and acetone carboxylase-coupled enzyme systems, as shown in

WO 03/039483 PCI7US02/36028

Figure 1 . Alternatively, the reaction could be monitored by following O2 consumption
electrochemically, or monitored optically by measuring absorbance or fluorescence of
NAD(P)H consumption as described below.

5 For other bacteria, including both aerobes and anaerobes, acetone metabolism is has been
identified as proceeding by a CC>2-dependent carboxylation reaction producing acetoacetate.
Acetone carboxylase, the enzyme that catalyzes this reaction, has recently been purified to
homogeneity from two bacterial sources. Although acetone carboxylase does not catalyze a
reaction that is readily detectable electrochemically, this enzyme has high specificity for
10 acetone and, according to the present invention, can be coupled with other enzymes that
catalyze redox reactions (for example dehydrogenases, oxidases). The feasibility of using
coupling enzymes with acetone carboxylase for electrochemical detection had not been
reported prior to this disclosure.

15 Suitable acetone carboxylases for use in an acetone-specific enzyme system include, but are
not limited to, acetone carboxylase obtained: from Xanthobacter autotrophicus strain Py2
(referred to herein as X. autotrophicus Py2 or as X. autotrophicus st. Py2) (see Sluis, M.K.
and Ensign, S.A., Purification and characterization of acetone carboxylase from
Xanthobacter strain Py2, PNAS USA, 94: 8456-8462 (1997)); from Rhodobacter capsulatus

20 BIO (see Sluis, M.K. et al., Biochemical, Molecular, and Genetic Analyses of the Acetone
Carboxylases from Xanthobacter autotrophicus Strain Py2 and Rhodobacter capsulatus
Strain BIO, /. Bacterial, 184(1 1):2969-77 (2002)); and from Rhodococcus rhodochrous
B276 (see Clark, D.D. and Ensign, S.A., Evidence for an inducible nucleotide-dependent
acetone carboxylase in Rhodococcus rhodochrous B276, J. Bact. 181(9):2752-58 (1999)).

25 Xanthobacter autotrophicus strain Py2 was deposited in the American Type Culture
Collection (ATCC) on October 29, 2002 under ATCC Accession No. PTA-4779. The

ATCC is located at 10801 University Boulevard, Manassas, VA 201 10-2209 U.S.A. and

may be contacted at P.O. Box 1549, Manassas, VA 20108 U.S.A. This deposit was made
in accordance with the requirements of the Budapest Treaty. The amino acid sequences of
30 the subunits of theX autotrophicus Py2 acetone carboxylase are set forth in SEQ ID NOs: 1,
2, and 3; the nucleotide sequences of the genes encoding these subunits are available in
GenBank (See accession number AY055852). The amino acid sequences of the
Rhodobacter capsulatus B10 acetone carboxylase gene are set forth in SEQ ID NOs:4, 5,

WO 03/039483 PCT/US02/36028

and 6; the nucleotide sequences of the genes encoding these subunits are available on the
website of the "Rhodobacter Capsulapedia" sequencing project (See
http://rhodol.uchicago.edu). Both theZ autotrophicus Py2 and the/?, capsulatus BIO
acetone carboxylase enzymes are alpha/beta/gamma (a/p/y) heterotrimers, sharing
approximately an 80% overall sequence identity with each other, as well as exhibiting
functional identity in catalyzing the same reaction with acetone.

Acetone-Specific Enzyme Systems

In a preferred embodiment of the invention, a breath acetone diagnostic device is provided
that contains one or more acetone-specific enzyme systems. A preferred use of such a
device is in monitoring ketone production in a mammal. In developing the invention, a
number of oxidoreductase enzyme systems were investigated that, in the presence of
acetone, oxidized pyridine nucleotides as cofactors or produced hydrogen peroxide as a co-
product, allowing the reaction be detected electrochemically. These oxidoreductase enzyme
systems include, for example: 1) the secondary alcohol dehydrogenase (S-ADH)-cataIyzed
reduction of acetone with concomitant NADPH consumption; 2) S-ADH-catalyzed
reduction of acetone with concomitant NADH consumption; 3) acetone carboxylase
reaction coupled to P-hydroxybutyrate dehydrogenase consumption of NADPH; 4) acetone
carboxylase reaction coupled to p-hydroxybutyrate dehydrogenase consumption of NADH;
5) acetone carboxylase reaction ATP hydrolysis coupled to NADPH consumption; 6)
acetone carboxylase reaction ATP hydrolysis coupled to NADH consumption; 7) S-ADH
reaction NADP + formation coupled to H 2 0 2 formation; 8) S-ADH reaction NAD + formation
coupled to H 2 0 2 formation; 9) acetone carboxylase reaction ATP hydrolysis coupled to .
H 2 0 2 formation; 10) acetone carboxylase reaction coupled to P-hydroxybutyrate
dehydrogenase NADP + formation coupled to H 2 0 2 formation; 1 1) acetone carboxylase
reaction coupled to P-hydroxybutyrate dehydrogenase NAD + formation coupled to H 2 0 2
formation; 12) acetone mono-oxygenase coupled to NADPH oxidation; 13) acetone mono-
oxygenase coupled to NADH oxidation; 14) acetone mono-oxygenase coupled to H 2 C>2
formation; and 15) acetone monooxygenase-catalyzed NAD(P)+ formation coupled to
H202 formation.

WO 03/039483 PCT/US02/36028

In all of these enzyme systems, the pyridine nucleotide or hydrogen peroxide is detectable
electrochemically, though other detection means known in the art can be utilized.

The use of enzymes as bioactive interfaces is well known in the art, and such interfaces are
5 used in analytical methods of detecting electronic transduction of enzyme-substrate
reactions. Direct electrical activation of enzymes such as redox enzymes permits
stimulation of bioelectrocatalyzed oxidation or reduction of enzyme substrates. Rapid
transfer of electrons between an electrode and a given redox enzyme results in current
generation corresponding to the rate of turnover of the electron exchange between the

10 substrate and biocatalyst. In other words, the transduced current of the system correlates
with enzyme substrate concentration. Electrical contacting of redox proteins in a biosensor
and the electrode support contained therein may be mediated by direct electron transfer with
electrode surfaces. Redox enzymes lacking direct electrical communication with electrodes
may achieve electrical contact by mediated electron transfer via active charge carriers. An

15 electron relay may be oxidized or reduced at an electrode surface, and diffusion of the
oxidized or reduced relay into enzyme results in short electron transfer distances with
respect to the active redox center for mediated electron transfer and, thus, electrical
activation of a biocatalyst.

20 Detection Means

The acetone-selective enzyme system, in acting upon the acetone substrate, generates an
electrochemically or non-electrochemically detectable product or by-product directly, or the
enzyme system will also include at least one further component. The further component
may be: one or more additional enzyme(s) forming an enzymatic pathway utilizing the

25 product or by-product of the initial enzymatic acetone reaction to thereby generate a

photometrically or electrochemically detectable product or by-product; or at least one signal
mediator; or both the additional enzyme(s) and the signal mediator(s). The signal
mediator(s) may be selected from, for example: indicators, such as a pH-change indicators;
electron transfer mediators; photometric mediators, and other components.

In an electrochemical embodiment of the invention, an acetone-specific redox enzyme or
enzyme system is selected that utilizes an electrochemically detectable cofactor, such as
N ADH, or generates a by-product, such as H2O2, during the course of the enzymatic

WO 03/039483 PCT/US02/36028
reaction. These enzyme systems can selectively detect acetone in biological samples, such
as breath or biological fluids. However, detection of acetone is not limited to
electrochemical means, and the enzyme system of this invention may be used in other types
of devices, for example devices employing known UV, fluorescence, or other suitable
5 methods of detecting acetone-specific enzyme-substrate interactions.

Non-Electrochemical Detection Means

Non-electrochemical detection involves, for example, any calorimetric or photometric
detection mode known in the art (for example, any colorimetric, spectrometry,
10 spectrophotometry, luminescence-based, chemiluminescence-based, or fluorescence-based
detection method.)

A fluorescence detection device has the following minimum requirements: it must be light-
tight to eliminate stray light from its surroundings, its fluors must be stored in the dark to

15 prevent photobleaching (that is increase shelf life), and its optics must be at a 90° angle. A
diode emitting the desired excitation wavelength can function as the light source, and a
PMT can function as the detector. These need not be elaborate since both the excitation and
emission of the fluor are known, and these are the only wavelengths required. The
same breath collection and acetone partitioning apparatus used in an enzyme

20 electrochemical device can be used in a fluorescence device. A portable fluorescence

detector for aflatoxin has been described in the literature (MA Carlson et a/., An automated
handheld biosensor for aflatoxin, Biosens. Bioelectr. 14:841 (2000)), so a precedent for a
portable fluorescence detector exists.

25 Both direct and indirect fluorescence allows the detection of acetone from both breath and
body fluids. The acetone-specific enzymes and their cofactors can be immobilized on a
disposable strip using conventional entrapment techniques. When acetone diffuses through
the immobilization medium to the enzyme, the acetone will be chemically altered.
Unfortunately acetone itself is not fluorescent and cannot be derivatized inside the

30 detection device. Thus another reagent needs to be derivatized with a fluorophore or a fluor
needs to be added to the system to monitor the reaction. For the secondary alcohol
dehydrogenase system, NADH consumption can be monitored, while the acetone
carboxylase system can use ATP-analogs. As the NADH or ATP-analog is consumed,

WO 03/039483 PCT/US02/36028
fluorescence intensity should decrease. Since the reaction with acetone is stoichiometric,
fluorescence intensity is proportional to acetone concentration. The H 2 02-generating
systems can use H2O2 and an additional fluor. In these systems, H2O2 production causes an
increase in fluorescence intensity that is proportional to acetone concentration.

We have verified that NADH in 100 mM phosphate buffer, pH 7.6, emits light directly at
approximately 470 nm when excited with 342 run light; these data agree with those reported
in the literature (MA Carlson et aL, 2000). In addition, NADH direct fluorescence has a
0.1-10 fiM linear working range, is independent of pH from pH 6-13, decreases in intensity
1.6% per °C, and exhibits little altered fluorescence intensity in the presence of cations and
enzymes below pH 10 (See PW Carr & LD Bowers, Immobilized Enzymes in Analytical
and Clinical Chemistry, In Chemical Analysis. A Series of Monographs on Analytical
Chemistry and Its Applications (PJ Elving & JD Winefordner, eds.; vol. 56, p. 122 (Wiley-
Interscience, New York, 1980), and references contained therein). Several groups have
described the use of direct NADH fluorescence to monitor enzymatic activity (AK Williams
& JT Hupp, Sol-gel encapsulated alcohol dehydrogenase as a versatile, environmentally
stabilized sensor for alcohols and aldehydes, 1 Am. Chem. Soc. 1998, 120:4366; and VP
Iordanov et aL, Silicon thin-film UV filter for NADH fluorescence analysis, Sens, Actuat. A,
2002, 97-98:161).

Indirect fluorescence of NADH can be detected using the dye rhodamine 123. Non-
radiative energy transfer (also called fluorescence resonance energy transfer, FRET) occurs
between the excited states of NADH and rhodamine 123. FRET is a well-known technique
for determining the proximity of two species, i.e. FRET is utilized as a "molecular
yardstick" both in vitro and in vivo. In this context of an acetone-specific enzyme system, a
donor fluorophore, e.g., NADH, transfers its excited state energies to the acceptor
fluorophore, rhodamine 123. (RP Haugland, Handbook of Fluorescent Probes and Research
Products. 2002 (9 th ed.; Molecular Probes, Inc.; Eugene, Oregon); K Van Dyke et aL, eds.
Luminescence Biotechnology. Instruments and Applications. 2002 (CRC Press; Boca
Raton, Florida) and references contained therein). The NADH-rhodamine 123 FRET
method has been successfully employed in other enzymatic assays (MH Gschwend et aL,
Optical detection of mitochondrial NADH content in intact human myotubes, Cell. MoL
Biol. 47:OL95 (2001); H. Schneckenberger et aL, Time-gated microscopic imaging and

WO 03/039483 PCT7US02/36028
spectroscopy in medical diagnosis and photobiology, Opt. Eng. 33:2600 (1994)).
Bioluminescence resonance energy transfer, or BRET, may also be used in conjunction with
an acetone-specific enzyme system according to the present invention. In BRET, the donor
fluorophore is replaced by a luciferase. Bioluminescence from luciferase in the presence of
5 a substrate excites the acceptor fluorophore. BRET has also been applied
in vitro and in vivo (K Van Dyke et al , 2002).

ATP can be derivatized with a fluorophore for indirect fluorescence. Several commercially
available dyes include BODIPY ATP and trinitrophenyl ATP (Haugland, 2002). These
10 analogs change their fluorescence intensity or become fluorescent when bound to an
enzyme's ATP binding site.

Indirect fluorescence detection of H 2 0 2 has also been reported (Carr & Bowers, 1980).
These methods utilize dyes that reduce the peroxide to H2O and are themselves oxidized.
15 Homo vanillic acid (4-hydroxy-3-phenylacetic acid) and /?-hydroxyphenylacetic acid are
among the most commonly used in clinical chemistry (Carr and Bowers, 1980). A
commercially available kit uses the dye Amplex Red for fluorescence detection of H 2 0 2
(Haugland, 2002).

20 Any fluorescent dyes and fluorescence-detectable enzyme substrate or cofactor analogs can
be used in a fluorescence device to detect acetone in breath or bodily fluids.

Chemiluminescence (CL) and electrogenerated chemiluminescence (ECL) (collectively
referred to herein as "(E)CL") are widely used in medical diagnostics and analytical

25 chemistry (C Dodeigne et al y Chemiluminescence as a diagnostic tool: A review, Talanta
2000, 51 :415; KA Fahnrich et al, Recent applications of electrogenerated
chemiluminescence in chemical analysis, Talanta 2001, 54:531). Enzyme-based (E)CL
systems are sensitive and specific, and many CL systems are used with enzyme cycling to
detect H 2 0 2 (Dodeigne et al, 2000). (E)CL can detect picomolar (pM; 1 0" 12 M)

30 concentrations of analyte over a wide linear range (Dodeigne et al, 2000; Fahnrich et al,
2001). An (E)CL device can be constructed in accordance with the following principles.
Since the reaction itself emits light, an (E)CL device does not need a light source. A
photomultiplier tube (PMT) can function as the detector; (E)CL is visible to the unaided,

WO 03/039483 PCT/US02/36028
dark-adapted eye. A battery can be the power source for ECL. ECL requires electrodes and
a source of applied potential. Like a fluorescence detection device, (E)CL devices need to
be light tight and their reagents need to be protected from light until use. Also like
fluorescence, (E)CL requires derivatized reagents or additional enzymes and reagents to
5 detect acetone. (E)CL devices can be used with disposable strips (BD Leca et al, Screen-
printed electrodes as disposable or reusable optical devices for luminol
electrochemiluminescence, Sens. Actual. B. 2001, 74: 190) and can be miniaturized (Y Lv et
al, Chemiluminescence biosensor chip based on a microreactor using carrier airflow for
determination of uric acid in human serum, Analyst 2002, 127:11 76).
10 An optical electrode (or optrode) can be fabricated using an acetone-specific enzyme system
according to the present invention. For example, an optrode such as that used in a glucose
optrode that uses ECL, may be employed (see CH Wang et al , Co- immobilization of
polymeric luminol, iron(II) tris(5-aminophenanthroline) and glucose oxidase at an electrode
surface, and its application as a glucose optrode, Analyst 2002, 1 27: 1 507)).

The most common CL systems involve the detection of H2O2 or another reactive oxygen
species (Carr & Bowers, 1980; Haugland, 2002; Dodeigne et al. 9 2000; K Van Dyke et al.,
2002) and references contained therein). The classic system is luminol-peroxidase. In basic
solution, H2O2 oxidizes luminol to an excited amino-phthalate ion; the excited amino-

20 phthalate ion emits a 425-nm photon to return to its ground state. When used in medical
diagnostics, this reaction is catalyzed with horseradish peroxidase (HRP) (Carr & Bowers,
1 980; Dodeigne et al., 2000). Thus any enzyme system that produces H2O2 or requires a
cofactor that can react with additional reagents to form H2O2 can be used in a CL device.
The H 2 C>2-generating systems described herein can use luminol-HRP directly for acetone

25 detection. These enzyme cycling schemes increase the light emission over time because the
substrates are continuously recycled (Dodeigne et al, 2000). While luminol itself is
frequently used in CL, its improved analogs can also be used in a CL-based detector
according to the present invention, in place of luminol, in order to increase the sensitivity.
Examples of such analogs are those described in Carr & Bowers, 1980; and Dodeigne et al,

30 2000.

NADH detection using CL is a common technique (Dodeigne et al, 2000). For example, in
the presence of l-methoxy-5-methylphenazinium methylsulfate, NADH reduces O2 to H2O2

WO 03/039483 PCT/US02/36028
which generates light using the Iuminol-peroxidase system (Dodeigne et ai 9 2000). For an
acetone monitor, the O2 in ambient air is sufficient to detect acetone using this system.
NADH also reacts with oxidized methylene blue to form H 2 0 2 that reacts with luminol
(Carr and Bowers, 1980). NADH can also act as a CL quencher. The fluorescence intensity
5 of the substrate ALPDO is decreased in the presence of NADH and HRP (Van Dyke et aL,
2002). NADH also can be used with Ru(bpy) 3 2+ for ECL (ES Jin et aL 9 An electrogenerated
chemiluminescence imaging fiber electrode chemical sensor for NADH, Electroanal 2001,
13(15):1287). Rhodamine B isothiocyanate can also be used for ECL detection of H2O2
(Fahnrich et al., 2001). ECL also offers another advantage in that, by use of a properly
10 poised electrode, the electroactive species can be regenerated at the electrode surface.

Regeneration both conserves reagents and allows durable and/or "reagentless' sensors. All
these systems can be used in a (E)CL device interfaced to an acetone-specific enzyme
system according to the present invention.

15 CL is widely used to quantitate ATP simply and sensitively (Carr & Bowers, 1980). The
enzyme luciferase catalyzes the reaction of ATP and luciferin to produce excited-state
oxyluciferin, which returns to its ground state with the emission of a 562-nm photon (Carr
& Bowers, 1980; Haugland, 2002). The quantum yield for this reaction is very high; 10" 14
mol ATP can be detected. A kit for this reaction is commercially available (Haugland,

20 2002). Because luciferase is the enzyme that causes fireflies to "glow," this reaction is
referred to as bioluminescence. Both native and recombinant luciferase are commercially
available, and several groups have reported using bioluminescence ATP assays to quantify
biological analytes (P Willemsen et al f Use of specific bioluminescence cell lines for the
detection of steroid hormone [antagonists in meat producing animals, Anal. Chim. Acta

25 2002, 473 : 1 1 9; S J Dexter et al , Development of a bioluminescent ATP assay to quantify
mammalian and bacterial cell number from a mixed population, Biomat. 2003, 24:nb27). In
addition to the luminol-HRP system, H2O2 can also be detected using peroxyoxalic acid
derivatives (Dodeigne et al. 9 2000). H 2 0 2 can also be detected with CL non-enzymatically
with ferricyanide as the catalyst (Dodeigne et al., 2000). In these (E)CL systems, the

30 acetone-specific enzymes described herein either produce H 2 0 2 or require cofactors that can
be utilized to form H2O2.

WO 03/039483 PCT/US02/36028

Optical biosensors use photometric detection (that is, absorbance, fluorescence) of
substrates consumed or products formed by the reaction catalyzed by the enzyme system
incorporated into the sensor. The acetone-specific enzyme reactions described may be .
monitored by several photometric methods-namely by measuring NAD(P)H absorbance at
5 340 nm for the pyridine nucleotide-dependent enzymes or absorbance of the quinoneimine
dye for the H2O2 forming enzyme systems. For the later, addition of a peroxidase allows
detection of H2O2 by catalyzing the reduction of H2O2 with concomitant oxidation of a dye
compound that upon oxidation absorbs at a specified wavelength. Peroxidase enzymes (for
example, commercially available horseradish peroxidase) typically have broad substrate
10 specificities so several different electron donor compounds may be used. NAD(P)H

consumption may also be measured by fluorescence detection (excitation at 350 nm and
emission at 450 nm).

Calorimetry may be employed as a detection means in an acetone-specific sensor according
15 to the present invention. Chemical reactions are typically either exo- or endothermic; that
is, they release or absorb heat as they occur. Calorimeters detect and measure this heat by
measuring a change in the temperature of the reaction medium (K Ramanathan & B
Danielsson, Principles and applications of thermal biosensors, Biosens Bioelectr. 16:417
(2001); B Danielsson, Enzyme Thermistor Devices. In Biosensor Principles and
20 Applications. Vol. 15, pp. 83-105 (LJ Blum & PR Coulet, eds.; Bioprocess Technology
Series, volume 15; Marcel Dekker, Inc: New York, 1991, pp. 83-105, and references
contained therein). Thus, the action of an acetone-specific enzyme or enzyme system may
be monitored calorimetrically. Calorimeters have been designed that are sensitive enough
to detect protein conformational changes, and calorimetry has been used to study many
25 enzymatic reactions in detail (M.J. Todd & J Gomez, Enzyme kinetics determined using
calorimetry: a general assay for enzyme activity? Anal. Biochem. 2001, 296:179 (2001)).

The major advantage of calorimetry is the lack of derivatization required for analysis
(Danielsson, 1991). Since most reactions involve heat exchange, and this heat is detected,
30 no chromophores, fluorophores, luminophores, "mediators," or other modifications of the
analyte are required. Reagents and analytes can be used "as is." This allows the analysis of
both reactions that lack a chromophore or fluorophore and/or would be difficult or
impossible to derivatize or couple to the generation of an electroactive species.

WO 03/039483

PCT/US02/36028

Miniaturized or chip-based thermosensors have been reported in the literature (Ramanathan
& Danielsson, 2001 ; B Xie & B Danielsson, Development of a thermal micro-biosensor
fabricated on a silicon chip. Sens. Actuat. B 6:127 (1992); P Bataillard et ah, An integrated
silicon thermopile as biosensor for the thermal monitoring of glucose, urea, and penicillin.
Biosen. Bioelect. 8:89 (1993)). These devices range from radically arranged thermopiles on
freestanding membranes to groups of thermopiles constructed on silicon/glass
microchanneis. These devices have been used to detect specific, single enzymatic reactions
(Danielsson, 1991; Xie & Danielsson, 1992; Bataillard et al., 1993). Moreover, two groups
have reported thermosensors for glucose (B Xie et al., Fast determination of whole blood
glucose with a calorimetric micro-biosensor, Sens. Actuat. B 15-16:141 (1993); MJ
Muehlbauer et al., Model for a thermoelectric enzyme glucose sensor, Anal. Chem. 61 :77
(1 989); BC Towe & EJ Guilbeau, Designing Medical Devices, 1 998,
http://lsvl.la.asu.edu/asubiotech/slideshow/slidel9.html (accessed January 2002).
Preliminary experiments using a conventional calorimeter indicate that the secondary
alcohol dehydrogenase-acetone reaction is exothermic (data not shown).

For the acetone monitor described herein, the acetone-specific enzymes and their cofactors
can be immobilized on a thermopile via conventional entrapment methods. The enzymes
and reagents associated with the coupled electrochemical detection, electrochemical
mediators, and "photonic" mediators (luminophores) are unnecessary for calorimetry. The
reaction involving acetone can be monitored directly without modification or derivatization.
When acetone in the breath or fluid sample diffuses through the immobilization medium
and encounters the enzyme, the acetone will be chemically altered. This reaction will
generate or absorb heat, causing a temperature change. Comparison of this temperature
with that of a reference thermopile will quantify this heat; the measurement is differential.
The quantity of heat released or absorbed is proportional to the analyte concentration.-

For breath collection, partitioning the acetone from the gas phase to the liquid phase, that is,
condensation, is exothermic. The reference or dual thermopile can compensate for this heat.
Thus an enzyme calorimetric acetone monitor can use the same breath collection apparatus
as an enzyme electrochemical acetone monitor except for the addition of the dual
thermopile.

WO 03/039483 PCT/US02/36028

The entire enzyme calorimetric device needs to be sufficiently insulated to prevent heat
exchange with its surroundings. Except for electrochemical detection, other aspects of the
device, such as enzyme stability, specificity, device portability, etc., described in this .
5 document are the same as those for an enzyme electrochemical device.

Thus, useful methods for achieving signal transduction in biosensors according to the
present invention include not only electrochemical (amperometric or potentiometric), but
also optical or photometric (including colorimetry, fluorescence-based techniques, or
10 chemiluminescence-based techniques), and calorimetric means, all of which are useful in
application to acetone biosensor signal transduction.

Therefore, although electrochemical detection means are described and exemplified in detail
herein, the enzyme systems of the invention are not limited to use in biosensors employing

15 electrochemical detecting strategies. Other detection strategies may be suitably integrated
into a biosensor specific for acetone in biological samples. Photometric assays, such as
assays in which changes in the amount of light absorbed in a reaction solution over time
may be used. Likewise, assays in which changes in fluorescence or changes in sample
turbidity may be employed for detecting acetone-specific enzyme-substrate interactions.

20 Such photometric assays are discussed hereinbelow. Redox potentials of H2O2 and

colorimetric/photometric detection of coenzymes is discussed by Bergmeyer. Photometric
assays for enzymatic activity are generally described by John in 'Thotometric Assays". An
NADH-consumption measuring electrode is disclosed by Hart et al. in a 1999 article
published in Electroanalysis. Vanysek discloses redox potentials in general, and oxidation-

25 reduction potentials of various compounds suitable for use in biochemical applications are
disclosed by Voet & Voet.

An enzyme system employing S-ADH coupled to alanine dehydrogenase was successfully
monitored spectrophotometrically for NADH formation. In addition, since the reaction also
30 generates ammonium ion, an optical sensor for NH/ can be employed as the detection means for
such an enzyme system. One such optical means is described by TD Rhines and MA Arnold,
Fiber-optic biosensor for urea based on sensing of ammonia gas, Anal Chim. Acta, 1989, 227:387;
several enzyme-based amperometric NH4 + sensors are commercially available. For acetone

WO 03/039483 PCT/US02/36028
detection, ammonia production can be coupled to the secondary alcohol dehydrogenase system
above; ammonia concentration would then be proportional to acetone concentration. Another
enzyme scheme to couple acetone to ammonia production is the following:

acetone + NADH + H* -> 2-propanol + NAD* (S-ADH)

glutamate + NAD + + H 2 0 a-ketoglutarate + NH/ + NADH (glutamate dehydrogenase)

This second scheme can be used either optically or amperometrically to detect acetone.
Additionally, the NADH is recycled. Likewise, an enzyme system in which acetone
carboxylase is coupled to glutamate dehydrogenase, generates NHL* 4 " and so can be detected
optically or amperometrically and correlated with acetone concentration.

Electrochemical Detection Means

Amperometric biosensors work by generating current between two electrodes by
enzymatically producing or consuming a redox-active compound. Several examples of
amperometric acetone biosensor schemes have been described in which NAD(P)H or H 2 0 2
are consumed or generated enzymatically in response to the presence of acetone. In
examples where the transducer is H 2 0 2 , an alternative means to monitor the reaction
amperometrically could be to employ a Clark-type oxygen electrode and measure a decrease
in 0 2 concentration. For example, in the case for the secondary alcohol dehydrogenase (S-
ADH) coupled to H 2 0 2 formation, the enzyme system catalyzes the following:

acetone + NADH + H + -> 2-propanol + NAD + (S-ADH)

lactate + NAD + -> pyruvate + NADH + H + (lactate dehydrogenase)

pyruvate + Pi + 0 2 ~> acetylphosphate + C0 2 + H 2 0 2 (pyruvate oxidase)

Oxygen is then reduced/consumed at the cathode generating a concentration gradient
between the electrode and the bulk solution. The rate of electrochemical reaction is
dependent on the oxygen concentration in solution.

Potentiometric biosensors employ ion-selective electrodes in which the release or
consumption of ions during an enzyme reaction is measured (for example, H + , CN\ NH 4 + )
(1,2, 3). For example, a potentiometric biosensor for measuring acetone concentration can

WO 03/039483 PCT/US02/36028

be utilized where NrL» + formation is coupled to the reaction catalyzed by S-ADH and
alanine dehydrogenase as follows:

acetone + NADH + H* 2-propanol + NAD + (S-ADH)
5 alanine + NAD + pyruvate + NADH + H* + NH 4 + (alanine dehydrogenase)

(Photometric data for this system has already been obtained to verify its utility for acetone-
specific signal transduction: see the discussion under the "Results" section, below).

10 A very similar system can be utilized with acetone carboxylase ( M p-OH-butyrate dehydr."
being "beta-hydroxbutyrate dehydrogenase"):

acetone + ATP + C0 2 -»acetoacetate + AMP + 2?i (acetone carboxylase)

acetoacetate + NADH + H* ->p-hydroxybutyrate + NAD + (p-OH-butyrate dehydr.)
1 5 alanine + N AD + ->pyruvate + NADH + H*+ NH/ (alanine dehydrogenase)

Another type of electrochemical biosensor that may be employed is a light-addressable
potentiometric sensor. In one embodiment of such a device, the acetone-specific enzyme
system(s) may be applied to (e.g., immobilized to the surface of) a potentiometric sensing
20 means such as that described, for sensing glucose, in A Seki et al., Biosensors based on
light-addressable potentiometric sensors for urea, penicillin, and.glucose, Anal Chim. Acta
373(l):9-13(2Nov. 1998).

In designing an acetone-specific biosensor according to the invention, various enzymatic
25 by-products and/or factors may be employed for the production of electrochemical signals.
One group includes organic cofactors, such as NAD, NADH, NADP, NADPH, FAD,
FADH, FMN, FMNH, Coenzyme A, Coenzyme Q, TTQ (Tryptophan Tryptophylquinone)
and PQQ (Pyrroloquinolinequinone). For example, a PQQ-dependent dehydrogenase may
oxidize isopropanol. Electrons from this reaction may be transferred through PQQ, which is
30 reduced, and can be oxidized at the electrode or with an intervening enzyme. Other
vitamins may also be used.

• 42

WO 03/039483 PCT/US02/36028

Enzymatic reaction by-products useful in the invention include hydrogen peroxide and
ammonium.

Energetic molecules may also be used in the invention for coupling acetone metabolism to
5 electrochemically measurable signals, including: ATP, ADP, AMP, GTP, GDP and GMP.
Neither these molecules nor phosphate can be detected directly, but can be detected through
coupling to a redox-by-product-producing enzyme system.

These by-products, cofactors, and energetic molecules can also be detected by non-
10 electrochemical means as described above.

Signal Mediators

Electron transfer mediators are redox-reversible species that may be used to transfer
electrons between (that is to or from) the electrically potentiated surface of an electrode and

15 an organic species (such as a co-factor) involved in an enzymatically catalyzed reaction.
Examples of electron transfer mediators include: ferrocene and derivatives, ferricyanide,
hydroquinone, benzoquinone and derivatives, 2,6-dichloroindophenol, methylene blue,
phenylenediamine and derivatives, phenoxazine and derivatives (for example, Meldola's
blue, that is 8-dimethylamino-2,3-benzophenoxazine), and phenazine alkosulfates (for

20 example, phenazine methosulfate, phenazine ethosulfate). In a given embodiment, one or
more than one species of electron transfer mediator may be used.

Electron transfer mediators can be used to improve the kinetics of electron transfer in a
given enzyme-coupled electrode system, since organic cofactors may easily impair detector

25 functions. This impairment is caused by the creation of free radicals via singly transferring
multiple electrons between organic species and the electrically potentiated surface of the
electrode. These free radicals then can exhibit dimer and/or polymer formation at the
electrically potentiated surface, which fouls the surface of the electrode, thereby inhibiting
efficient electron transfer. Electron transfer mediators can be employed to avoid this

30 fouling of electrodes. Electron transfer mediators may also be used in situations where a
shift in electrode voltage is desired, for example, where the preferred voltage for use in the
reaction system without such a mediator happens to be a potential at which too much
electrical interference ("noise") occurs. An electron transfer mediator may be added in

WO 03/039483 PCT/US02/36028

order to permit a shift in the applied voltage to a different voltage region in which less noise
occurs. Examples of diffusional electron-transfer mediators applicable to immobilized
enzymes such as glucose oxidase, horseradish peroxidase, and the like, are set forth in Table
SofWillner and Katz.

Preferred mediators useful in multi-electron transfers for reduced forms of, for example,
NADH, NADPH, FADH, FMNH, Co-Q, PQQ, include, for example: ferrocene and
derivatives, ferricyanide, hydroquinone, benzoquinone and derivatives, 2,6-
dichloroindophenol, methylene blue, phenylenediamine and derivatives, phenoxazine and
10 derivatives (for example, Meldola's blue, that is 8-dimethylamino-2,3-benzophenoxazine),
and phenazine alkosulfates (for example, phenazine methosulfate, phenazine ethosulfate).

A second group of mediator factors that may be employed for the production of
electrochemical signals include inorganic cofactors such as Pt, Os, V, Mn, Fe, Co, Ni, Cu,
15 Mo, and W (see Holm et al., Aspects of Metal Sites in Biology, Chem. Rev. 1996. 96,
p.2239-2314). Some useful enzymes contain a heme center, and thus iron (for example,
cytochrome P450 monooxygenase). Also useful is amine oxidase, which contains Cu.

In an alternative embodiment, a "photometric mediator" may be added to the enzyme
20 system in order to react with a product or by-product of the enzymatic reaction(s) and
thereby generate a derivative that can be, for example, photochemically, colorimetrically,
fluorometrically, or (UV or IR) spectrometrically detected. Thus, the addition of such a
"photometric mediator" may be characterized as permitting the conversion of a result of the
enzymatic reaction, that is a product or by-product, into a photometric signal. For example,
25 in the case of enzymatically catalyzed redox reactions, a chromogenic redox indicator such
as, for example, a tetrazolium salt, may be used as the photometric mediator. Many such
chromogenic redox indicators are known in the art. Examples of tetrazolium salts include,
but are not limited to: 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-2H-tetrazolium bromide
(MTT bromide); (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-
30 sulfophenyl)-2H- tetrazolium, inner salt (MTS; available from Promega Corp., Madison,
Wisconsin); and (5-cyano-2,3-ditolyl tetrazolium chloride) (CTC). Such photometric
mediators can be used, for example, to convert the redox "signal" of an electron transfer
mediator into a photometrically detectable signal.

WO 03/039483

PCT/US02/36028

The enzymes and other components of the enzyme system may be immobilized in a gel
layer disposed upon the electrode surface. Any of the various gels known in the art as
useful for immobilization of biologies in the presence of an electrode may be used. For
example, a method such as is described in PCT/US02/16140 (filed May 21, 2002) may be
used to immobilize the biologic components of an acetone-specific enzyme system in a
polyurethane hydrogel disposed upon an electrode.

Enzyme Arrays and Multi-Enzyme Systems

In addition to monitoring acetone in a sample per se, acetone-specific enzyme systems may
be useful in biosensors having more than one enzyme system for detecting multiple
substrates in a given biological sample. For example, an electrode linked to an acetone-
specific enzyme system may enable subtraction of an acetone signal from an ethanol
detector. Such a set up could be configured in an array, wherein at least two different
detection modes or at least two different detectors would be operative for detecting ethanol
and acetone. Such an array would be useful to correct for acetone interference in ethanol
breathalyzer analyses.

Fluorescence detection can also be accomplished using arrays. Fluorescence sensor arrays
have been described in the literature. They have been used for such complex samples as
wine aromas, perfume, and genes. Fluorescence sensor arrays employ fluorescent or
chromogenic dyes or substrates that covalently attached to polystyrene beads in wells on the
distal face of an optical fiber (D.R, Walt, Imaging optical sensor arrays. Curr. Opin. Chem.
Biol. 6:689 (2002)). A high-density optical array can contain several types of dyes or
substrates for different analytes. The array is exposed to each possible component
individually, then to the sample. Pattern recognition is employed to deduce the composition
of the sample. In the case of breath or bodily fluid components, the acetone-specific
enzymes, cofactors, and chromogenic or fluorogenic dyes can be covalently attached to a
portion of beads, while enzymes specific for other analytes, such as ethanol, can be attached
to other beads. Each bead will "light up" upon exposure to its target analyte.

An enzyme-based fluorescence or chromogenic array has never been applied to the
detection of acetone.

WO 03/039483 PCT/US02/36028
Uses for an Acetone-Specific Enzymatic Biosensor

Breath acetone monitoring is a useful tool for monitoring effectiveness and compliance of
subjects on weight loss diets. Ketosis can be manipulated by exercise and dieting choices,
5 even between two diets with equal energy balance. The response time for reflecting diet
and exercise choices in breath acetone levels is in the order of 2-3 hours, and was a better
indicator of fat loss than urine ketone analysis.

A home acetone diagnostic biosensor would be useful in aiding subject management of
Type 1 and Type 2 diabetes. Such biosensors would enable subjects to monitor weight loss,
to detect signs of the onset of ketoacidosis, and to control sugar intake with respect to
insulin availability, especially in Type 1 diabetics. Indicators suggest that weight loss
success would be improved if subjects could share daily acetone measurements with health
care professionals and peers via the Internet and weekly support group meetings. Use of the
inventive acetone-specific detection system is not limited to management of obesity and
diabetes. It is contemplated that the acetone-specific biosensors described herein would be
useful for managing any disease in which acetone production is an indicator of pathology.

In addition, the acetone-specific enzyme system may prove to be a highly effective means
of monitoring subject compliance with prescribed therapeutic regimes via drug tagging with
acetoacetate or a derivative thereof. The degradation of acetoacetate to acetone could be
measured via a biosensor containing the inventive acetone-specific enzyme system, thereby
improving the ability of health care professionals to track the dosing and bioavailability of
the corresponding tagged drug.

EXAMPLES

Acetone-specific enzyme systems and acetone sensors utilizing these systems have been
developed. Enzyme identification and/or purification, enzyme characterization and
30 selection, enzyme-plus-cofactor systems, multiple-enzyme-plus cofactor systems, coupled
enzyme systems providing linear (stoichiometric) acetone detection, coupled enzyme
systems providing amplified (for example, exponential) acetone detection, enzyme and
enzyme system stability testing, acetone vapor-to-liquid partitioning studies, and enzyme-

WO 03/039483 PCT/US 02/36028

mediated acetone sensor devices (both electrochemical and non-electrochemical devices)
that utilize such systems sensors are disclosed below in particular exemplified
embodiments. These examples are provided for exemplification and are not intended to
limit the invention. Particular embodiments employing acetone-specific enzyme systems in
enzyme-based electrochemical and non-electrochemical sensors is described below.

MATERIALS & METHODS

Materials. Acetone carboxylase from X autotrophic™ strain Py2 and isopropanol-grown X
autotrophics strain Py2 cell paste were obtained from Professor Scott A. Ensign at Utah
State University, Logan, UT. Acetone carboxylase from Rhodobacter capsulatus BIO
(ATCC 33303), acetone-grown R. capsulatus B10 cell paste, propane-grown
Mycobacterium vaccae JOB5 (ATCC 29678) cell-free extracts, and propane-grown
Rhodococcus rhodochrous B276 (ATCC 31338) cell paste were also obtained from
Professor Ensign, and all of these bacterial strains are publicly available. Secondary alcohol
dehydrogenase froral autotrophicus strain Py2 and isopropanol-grown X. autotrophics
strain Py2 were isolated from cell paste; and exemplary, publicly available secondary
alcohol dehydrogenases are described in Table 1. Pyruvate kinase (EC 2.7.1.40),
myokinase (EC 2.7.4.3), pyruvate oxidase (EC 1.2.3.3), horseradish peroxidase (EC
1.11.1.7), lactate dehydrogenase (EC 1 . 1 . 1 .28), malic enzyme (EC 1.1.1 .40), alcohol
dehydrogenase (EC 1.1.1.2), alanine dehydrogenase (EC 1.4.1.1), alcohol dehydrogenase
(EC 1.1.1.1), alcohol oxidase (EC 1.1.3.13), and P-hydroxybutyrate dehydrogenase (EC
1.1 .1.30) were purchased from Sigma (St. Louis, MO). All other chemicals and reagents
used were analytical grade. All solutions were prepared in 18 MQ water (Millipore).

WO 03/039483 PCT/US02/36028

TABLE 1. Information for Some Publicly Available S-ADH Enzymes

Cofactor

Organism

Source

Reference(s) [& Comments]

NADPH

Thprmnnnnprf)-

A fiCf ffHJUflLltJf KJ

bium brockii

Sipma Chem

Co. (catalog
no. A8435)

RJ I amed et al Enzvme & Microh

Technol., 3:144 (1981);

RJ Lamed & JG Zeikus, Biochem. J,

195(l):183-90 (1 Apr 1981);

A Ben-Bassat et al., J Bact, 146(l):192-99

(Apr 1981V 1

[DNA sequence available in GenBank (Acc.

No. X64841)]

NADH

A4vro bacterium
vaccae strain
JOB5 (Gram-
positive)

ATCC 29678

JP Coleman et al J Gen Microbiol

131(1 l):290l-07 (Nov 1985);
[Describes enzyme purification]

NADH

Pseudomonas sp.
6307 [CRL 75]
(Gram -negative)

ATCC 21439

CT Hou et ah, Eur. J Biochem., 1 19(2):359-
64 (Oct 1981);

IDe^cribes enzvme mirificationl

NADH

Xanthobacter
autotrophicus
strain Py2

ATCC PTA-
4779

[Enzyme purification and characterization
described herein]

NADPH

Thermoanaero-

hnctPT pthnnnlinj?

39E

ATCC 33223

DS Burdette et al., Biochem. J 3 1 6( 1 ): 1 1 5-22

[Describes enzyme purification, gene cloning
& DNA sequencing]

NADH

Candida utilis

DSM 70167;
ATCC 26387

H Schutte et al., Biochim. et Biophys. Acta,
716f3V298-307 (16 Jun 1982V

/ 1 Ul J / .Lf7D -J \J 1 1 1U J Ull 1 70Z. K

[Describes screening for S-ADH activity in
several yeast strains]

NADPH

"Anaerobic
extremely
thermophilic .
bacterium"

Biocatalvsts
Ltd. (Wales;
catalog no.
S300)

NADH

Candida boidinii
(yeast)

Fluka

(Milwaukee,
WT; catalog
no. 91031)

NADH

Candida sp.
(yeast)

NovaBiotec
Dr Fechter
GmbH (Berlin,
Germany;
catalog no.
"Isopropanol
dehydrogenase
(E.C.

1.1.1.80)")

WO 03/039483

PCT/US02/36028

Enrichment and isolation of acetone-, isopropanol-, and propane-utilizing microorganisms.
Enrichment cultures were set up in 160 mL serum bottles that were crimp-sealed with butyl
rubber stoppers. The bottles contained 10 mL mineral salts medium containing (in g/L):
NaNH4HP0 4 (1.74); NaH 2 P0 4 x H 2 0 (0.54); KC1 (0.04); MgS0 4 x 7 H 2 0 (0.2) and 1 mL/L
of a trace element stock solution (stock solution (in g/L): FeCl 2 x 4 H 2 0 (5.4); MnCl 2 x 4
H 2 0 (1.0); ZnS0 4 x 7 H 2 0 (1.45); CuS0 4 x 5 H 2 0 (0.25); concentrated HC1 (13 mL/L);
(NH4)6Mo 7 0 24 x 4 H 2 0 (0.1); H 3 B0 3 (0.1); CoCl 2 x 6 H 2 0 (0.19)). The pH of the medium
was adjusted to pH 7.2. Enrichments for propane-utilizing microorganisms were inoculated
with about 0.5 g of soil that had been purchased from a local supplier of top soils, or with
about 0.5 g of non-sterilized potting soil or organic compost that had been purchased from a
local supermarket.

Gaseous propane was added with a syringe to a 20% (v/v) concentration in the headspace of
the serum bottle. Enrichments for acetone- and isopropanol-utilizing microorganisms were
set up in a similar way except that substrates were added from a 1 M stock solution to a
final concentration of 25 mM acetone, or 10 mM isopropanol. The enrichment cultures
were incubated on a shaker at 28°C. For the isolation of single colonies, enrichment
cultures were cultivated on mineral salts medium (as described above) containing 1.5% w/v
agar (hereinafter "mineral salts agar").

In a different set-up, enrichment cultures were started for acetone-utilizing microorganisms
that could grow in the presence of a C0 2 -trap. 20 mL of mineral salts medium with trace
elements (see above) was filled into 250 mL baffled Erlenmeyer flasks. The medium was
inoculated with about 0.5 g of soil sample (see above). The Erlenmeyer flask was closed
with a rubber stopper that had been modified to hold a glass bulb. The glass bulb contained
about 0.5 mL of 50%(w/v) KOH. The KOH trapped the C0 2 from the Erlenmeyer flask
headspace. These set-ups were designed to enrich for acetone-utilizing microorganisms
with an acetone carboxylase-independent pathway. The enrichment cultures were incubated
on a shaker at 28°C.

Enrichment cultures were transferred two to three times after turbidity indicated bacterial
growth (usually after 3 to 5 days). To isolate single colonies, enrichment cultures were

WO 03/039483 PCT/US02/36028
spread on mineral salts agar plates. For the isolation of propane-utilizers, the agar plates
were placed in a 3.5 L anaerobic jar. Propane was added to the jar until a positive pressure
of 0.3-0.5 bar was reached inside the jar. The jar was placed into an incubator at 28°C. For
the isolation of acetone-utilizing microorganisms, the agar plates were placed into a 1 .4 L
5 desiccator. The desiccator contained two open glass vials with 3-4 mL neat acetone each.
The desiccator was sealed with several layers of PARAFILM (a wax-based sealing film,
from American National Can, Chicago, IL) before it was placed in an incubator at 28°C.
For the isolation of acetone-utilizing microorganisms that would grow in the presence of a
CCVtrap, agar plates were placed in a desiccator as described above. In addition to a vial

10 with acetone, a vial containing 50% KOH (about 4 mL) was placed into the desiccator.
Alternatively, for the isolation of acetone- and isopropanol-utilizers, enrichment cultures
were transferred to agar plates containing mineral salts medium plus acetone or isopropanol.
Additional acetone or isopropanol was added onto a small foam plug that was placed inside
the Petri dish. The Petri dish was sealed with several layers of parafilm to reduce

15 evaporation of substrates during incubation.

Colonies were visible on the agar plates after 5-10 days. Isolates were transferred to fresh
agar plates and incubated as described above. Isolated strains were also streaked onto
nutrient agar to check for purity. After several transfers on agar plates, 3 1 strains were
20 isolated that looked different as evaluated by colony morphology. Eight strains were

isolated from propane enrichments (these were designated TDCC Prop 1-8), eight strains
were isolated from isopropanol enrichments (these were designated TDCC IP- 1-8) and
fifteen strains were isolated from acetone enrichments (these were designated TDCC Ac 1 -
1 5). None of these were obtained from an acetone + KOH-trap enrichment.

Screening of isolates and culture collection strains for growth on acetone, propane, or
isopropanol. Isolates and culture collection strains were screened for growth on acetone,
propane, and isopropanol in 60 mL-serum vials containing 5 mL of medium plus 0.005%
(w/v) yeast extract as described above. The medium was inoculated from a single colony.
30 Isopropanol was added from a stock solution to a final concentration of 8 mM. Cultures
that showed more turbidity with substrates compared to cultures without substrates
(medium blanks) were considered hits. Hits were then screened for growth with acetone in
the presence of a C0 2 -trap as described above.

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PCT/US02/36028

Cultivation of isolates/strains, harvesting, and preparation of cell-free extracts. Several
isopropanal-utilizing strains were cultivated in larger batches for initial purification of
secondary alcohol dehydrogenase and experiments with cell-free extracts. Strain
5 Rhodococcus rhodochrous B276 (ATCC 31338) (formerly Nocardia corallina B276) and
strain TDCC IP-1 (and two additional strains: data not shown) were cultivated in 2 x 500
mL batches of mineral salts medium (for composition see above) plus 0.005% yeast extract.
Isopropanol (8 mM) was added initially as carbon and energy sources. Cultures were
incubated on a shaker (200 rpm) at 30°C. Growth was followed by monitoring the optical

10 density at 600 nm. More isopropanol was added at several time points when the growth rate
decreased due to lack of substrate. A total of about 96 mM isopropanol was added to the
cultures. At the end of the logarithmic growth phase, cells were precipitated by
centrifiigation (GSA rotor, 8,500 rpm, 20 min.) at 4°C. The cells were washed once in 50
mM Tris-HCI buffer, pH 7.5. The cell pellet was weighed and resuspended in a small

1 5 volume of TRIS (2-amino-2hydroxymethyM ,3 -propanediol) buffer (about 2 mL per g cells
(wet weight)). Cells were frozen at -20°C until further use. For the preparation of cell-free
extracts, cells were thawed and broken by sonication (4 x 20 s, pulsed, 50% intensity). Cell
debris and unbroken cells were precipitated by centrifiigation for 5 min. at 14,000 rpm (in
an Eppendorf benchtop centrifuge, Model 5417C, Brinkmann, Instruments, Inc., Westbury,

20 NY). Alternatively, for larger preparations, the cell suspension was passed three times
through a mini-French pressure cell at 20,000 psi (137,895.2 kPa), and the iysate was
clarified by centrifiigation at 6,000 x g for 40 min at 4°C.

Protein purifications. Acetone carboxylase from X autotrophicus strain Py2 and acetone
25 carboxylase from R. capsulatus were purified as described previously. Secondary alcohol
dehydrogenase (S-ADH) from X autotrophicus Py2 was purified via the following
protocol. Cell-free extracts (380 mL) of isopropanol-grown X autotrophicus Py2 (150 g)
were prepared as described above, and applied to a 5 x 15 cm column of DEAE-Sepharose
FAST FLOW (Diethylaminoethyl cross-linked agarose bead material; Catalog number 17-
30 0709- 1 0, Amersham Pharmacia Biotech, Piscataway, NJ)) equilibrated in buffer A (25 mM
MOPS (3-(N-morpholino)propanesulfonic acid), pH 7.6, 5% glycerol, ImM dithiothreitol)at
a flow rate of 10 mL/min. After loading, the column was washed with 1000 mL buffer A
and developed with a 2400 mL linear gradient of 90-290 mM KC1 in buffer A. Fractions

WO 03/039483 PCT/US02/36028
containing S-ADH activity were pooled and dialyzed against 2 L of 25 mM potassium
phosphate (pH 6.2) containing 5% glycerol (buffer B) for 16 h at 4°C. The protein was then
applied to a RED SEPHAROSE CL-6B (Procion Red HE-3B dye-linked, cross-linked-
agarose bead material, affinity matrix for affinity chromatograph; Catalog number 17-0528-
5 01 , Amersham Pharmacia Biotech) column (1.5x10 cm) equilibrated in buffer B at a flow
rate of 2 mL/min. After washing the column with 30 mL of buffer B, S-ADH was eluted
with 20 mL of buffer A containing 10 mM NAD + . Fractions containing S-ADH were then
dialyzed against 2 L of buffer A for 16 h at 4°C, concentrated by ultrafiltration (using a
YM30 ultrafiltration membrane; catalog no, 13722, from Millipore, Bedford,

10 Massachusetts), and frozen in liquid nitrogen. Partially purified S-ADH from bacterial
screen cultures was prepared as follows: cell-free extracts from 1 to 5 g of cell paste were
prepared as described above and applied to a 5 mL HI TRAP Q column (quaternary,
tetraethylammonium, cross-linked agarose bead material for use as an anion exchange
matrix; catalog number 17-1153-01, Amersham Pharmacia Biotech) equilibrated in 100 mM

15 MOPS, pH 7.6, containing 5% (v/v) glycerol (buffer C). The column was washed with 10
mL buffer C and developed with a 100 mL linear gradient of 0 to 100 mM NaCl in buffer C.
Fractions containing S-ADH activity were pooled, concentrated to 0.5 mL using a 30 kDa
MWCO ("molecular weight cut-off) centrifugal membrane (catalog number UFV4BTK25,
Millipore), and stored at -80°C.

Example 1

Acetone carboxylase coupled to NADH oxidation spectrophotometry assay.

Assays were performed in 2 mL (1 cm path length) quartz cuvettes that had been modified

by fusing a serum bottle-style quartz top (7x13 mm at mouth), allowing the cuvettes to be

25 sealed with a red rubber serum stopper. The reaction mix contained ATP (10 mM), MgCh
(1 1 mM), potassium acetate (80 mM), MOPS (100 mM), C0 2 (50 mM (1 mol C0 2 (g) to 4
mol potassium bicarbonate to maintain pH)), and 20 to 40 \ig purified acetone carboxylase
in a total volume of 1 mL at pH 7.6. The addition of P-hydroxybutyrate dehydrogenase (3
U) and NADH (0.2 mM) allowed acetoacetate formation to be coupled to the oxidation of

30 NADH. Assays were pre-incubated for 2 min. at 30°C with all assay components except
acetone. Assays were initiated by addition of acetone (2 mM). The reaction was monitored
by measuring the decrease in absorbance at 340 nm (e 3 4o of 6.22 mM^-cm' 1 for NADH)

WO 03/039483 PCT/USO 2/36028

over time in an Agilent Technologies (Palo Alto, CA) model 8453 UV-Visible
Spectroscopy System containing a thermostat-controlled cell holder at 30°C.

Acetone carboxylase coupled to H 2 0 2 formation spectrophotometry assay. Assays were
5 performed in 2 mL (1 cm path length) quartz cuvettes and contained ATP (0.1 mM), MgS(>4
(10 mM), potassium acetate (80 mM), potassium phosphate (50 mM), C0 2 (50 mM (1 mol
C02(g) to 4 mol potassium bicarbonate to maintain pH)), 40 ng purified acetone
carboxylase, phosphoenolpyruvate (2 mM), pyruvate kinase (20 U), myokinase (15 U),
pyruvate oxidase (2 U), peroxidase (15 U), flavin adenine dinucleotide (0.01 mM),

1 0 cocarboxylase (0.2 mM), 4-aminoantipyrine (0. 5 mM), and W-ethyl-iV-(2-hydroxy-3-
sulfopropyl)-m-toluidine (0.02% w/v) in a total volume of 1 mL at pH 7.5. Coupling
enzymes and reagents (that is phosphoenolpyruvate, pyruvate kinase, myokinase, pyruvate
oxidase, flavin adenine dinucleotide, and cocarboxylase,) allowed ATP hydrolysis to be
coupled to H2O2 formation (pyruvate oxidation). Addition of peroxidase, 4-

1 5 aminoantipyrine, and A^-ethyl-//-(2-hydroxy-3-sulfopropyl)-/n-toluidine allowed H 2 0 2
formation to be monitored spectrophotometrically at 550 nm (8550 of 36.88 mM" , -cm" 1 for
quinoneimine dye product) over time in a thermostat-controlled cell holder at 30°C. Assays
were pre-incubated for 2 min. at 30°C with all assay components except acetone. Assays
were initiated by addition of acetone (5 mM).

Example 2

Secondary alcohol dehydrogenase NADH oxidation spectrophotometry assay.
Assays were performed in 2 mL quartz cuvettes and contained NAD(H) (0.2 mM),
potassium phosphate buffer (25 mM), and a source of enzyme (cell-free extracts, column
25 fractions, or purified enzyme) in a total reaction volume of 1 mL at pH 6.2 (for ketone
reduction assays) or pH 7.8 (for alcohol oxidation assays) at 30°C. Assays were pre-
incubated for 1 .5 min. at 30°C with all assay components except substrate. Assays were
initiated by addition of substrate (2.5 mM) and monitored over time by measuring the
change in absorbance at 340 nm (e 3 40 of 6.22 mM* l 'cm' 1 for NADH).

Secondary alcohol dehydrogenase coupled to H 2 0 2 formation spectrophotometry assay.
Assays were performed in 2 mL (1 cm path length) quartz cuvettes and contained potassium
phosphate (50 mM), 1.5 jag purified S- ADH, NADH (50 \xM\ lactate ( 1 0 mM), lactate

WO 03/039483 PCT/US02/36028
dehydrogenase (20 U), pyruvate oxidase (2 U), peroxidase (15 U), flavin adenine
dinucleotide (0.01 mM), cocarboxylase (0.2 mM), 4-aminoantipyrine (0.5 mM), and N-
ethyI-A42-hydroxy-3-sulfopropyl)-m-toluidine (0.02% w/v) in a total volume of 1 mL at
pH 6.2. Coupling enzymes and reagents (that is lactate, lactate dehydrogenase, pyruvate
5 oxidase, flavin adenine dinucleotide, and cocarboxylase) allowed NADH oxidation to be
coupled to H2O2 formation (pyruvate oxidation). In some assays (where specified), lactate
and lactate dehydrogenase were replaced with alanine (10 mM) and alanine dehydrogenase
(2 U). Assays were monitored spectrophotometrically at 550 nm (e 55 o of 36.88 mM' , xm* 1
for quinoneimine dye product) over time in a thermostat-controlled cell holder at 30°C as
10 described above. Assays were pre-incubated for 2 min. at 30°C with all assay components
except acetone. Assays were initiated by addition of acetone (2.5 mM).

Primary alcohol dehydrogenase coupled to primary alcohol oxidase substrate recycling
assays.

15 Assays were performed in 2 ml (1 cm path length) quartz cuvettes and contained potassium
phosphate (25 mM), alcohol dehydrogenase (1 U), NADH (100 fiM), alcohol oxidase (2 U),
peroxidase (15 U), 4-aminoantipyrine (0.5 mM), and//-ethyl-7V-(2-hydroxy-3-sulfopropyl)-
m-toluidine (0.02% w/v) in a total volume of 1 mL at pH 6.2. Assays were monitored
spectrophotometrically at 550 nm (6550 of 36.88 mM^-cm" 1 for quinoneimine dye product)

20 or at 340 nm (e 34 o of 6.22 mM' , 'cm' 1 for NADH) over time in a thermostat-controlled cell
holder at 30°C as described above. Assays were pre-incubated for 2 min. at 30°C with all
assay components except ethanol. Assays were initiated by addition of ethanol (50 or 5
|iM).

25 Stability studies. A sufficient quantity of enzyme for each individual activity assay (for
example, 1.5 |ig S-ADH) was aliquoted into 1.5 mL microcentrifuge tubes with specified
concentrations of additives (for example trehalose (10% w/v)) in buffer (25 mM MOPS, pH
7.6) and frozen at -80°C for 1 h. Samples were then placed in a shelf freeze dryer (Virtis
model Advantage ES) and held at -50°C (shelf temperature) for 16 h, and then increased to

30 20°C for 4 h. Freeze-dried samples were removed and allowed to sit at room temperature
(1 7 to 24°C) over time. At specified time points, samples were re-hydrated and assayed as
described above.

WO 03/039483 PCT/US02/36028

Protein characterizations. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis
(SDS-PAGE) was performed following the Laemmli procedure (Laemmli, U.K., Nature,
227:680-685 (1970)) using a 12%T, 2.7%C gel. "%T" indicates weight percent of total
5 monomers, a measure of total monomer concentration, which is given by %T = 1 00 x
((grams acrylamide) + (grams cross-Iinker))/total gel volume (in mL); "%C" indicates
weight percent of cross-linker, which is given by %C = 100 x (grams cross-linker)/((grams
acrylamide) + (grams cross-linker)); and the cross-linker used was N,N'-methylene-bis-
acrylamide. Electrophoresed proteins were visualized by staining with Coomassie Blue

1 0 (PhastGel Blue R, catalog number 1 7-05 18-01, Amersham Pharmacia Biotech). The
apparent molecular masses of polypeptides based on SDS-PAGE migration were
determined by comparison with Rvalues of standard proteins. N-terminal sequencing was
performed by Commonwealth Biotechnologies, Inc. (Richmond, VA). Protein
concentrations were determined by using a modified biuret assay (V.J. Chromy et al., Clin.

15 Chem, 20:1362-63 (1974) with 5-globulin as the standard.

Mass spectrometry analysis of enriched S-ADH and generation of peptide amino acid
sequences was performed as follows. The S-ADH soluble protein was characterized by
high-resolution two-dimensional gel electrophoresis. Proteins (30 jig) were solubilized for

20 isoelectric focusing (IEF) analysis in rehydration sample buffer consisting of 5 M urea, 2 M
thiourea, 2% (w/v) CHAPS (3-[(3 -cholamidopropy l)dimethy lammonio]- 1 -
propanesulfonate), 2% (w/v) SB 3-10 (2-(decyldimethylammonio)propanesulfonate), 40
mM TRIS, 2 mM tributyl phosphine (added to rehydration solution just before use), and
0.2% Bio-Lyte 3/10 (Bio-Rad, Hercules, CA, cat no. 163-2104). Protein/rehydration

25 solution was rehydrated into 1 1 cm IPG ReadyStrip pH 3-10 (Bio-Rad, Cat. no. 163-2014)
under passive conditions 0 volts, 20 °C, 16 hrs.

One-dimensional isoelectric focusing was carried out on a Protean IEF cell (Bio-Rad, model
no. 526BR02142) for 35,000 volt-hours using IPG ReadyStrips (Bio-Rad). Following first
30* dimension electrophoresis, gels were equilibrated for 20 minutes in a buffer containing 20
% glycerol, 0.375 M Tris, 6 M urea, 2% SDS, and 5 M tributyl phosphine. IPG
ReadyStrips were placed on top of a Criterion™ precast 1 mm 4-20% gradient Tris-HCl-
SDS gel (Bio-Rad, cat. no. 345-0036) and 0.5% warm Agarose containing 0.01%

WO 03/039483 PCT/US02/36028

bromophenol blue (Bio-Rad, cat. no. 161-0404) was added to the remaining well.
Electrophoresis was carried out on a Criterion mini electrophoresis cell (Bio-Rad, cat. no.
165-6001) at room temperature. The electrophoresis running buffer was prepared from a
10X Tris-glycine-SDS solution (Bio-Rad, cat. no. 161-0732). Following assembly of the
5 gel system and addition of the running buffer, the electrophoresis was carried out at an
initial current of 2 mA, 3500 volts, 45 watts, for 1.5 hrs. The current was ramped up to 5
mA for 30 minutes followed by 10 mA for 2-3 hrs. Typical run times were between 4-5 hrs.
Following electrophoresis, gels were stained in a buffer consisting of 17% ammonium
sulfate, 30% methanol, 3% phosphoric acid, and 0.1% coomassie brilliant blue G250 (Bio-
10 Rad, cat no. 161-0436), for at least 12 hrs. Gels were rinsed with water and stored in 2%
acetic acid until further processing.

Colloidal Coomassie-stained gel images were captured using Bio-Rads Fluor-S
Multilmager (Bio-Rad, cat. no. 170-7700). Digital filtering algorithms were used to remove
15 non-uniform background without removing critical image data. Internal standards
(molecular weight markers) were used initially to determine the molecular weight of the
targeted proteins of interest. The molecular weight and pi of the S-ADH protein were
determined by comparison of its position on the two-dimensional gel relative to the protein
standards.

Protein spots relative to S-ADH from the 2-D gel were excised manually. The gel pieces
. . were macerated and destained with 25 mM ammonium bicarbonate/50% acetonitrile in a 1.5
mL microfuge tube with vigorous shaking for 30 minutes. The blue-tinted destaining
solution was removed and discarded with a fine-tip pipette. The destaining step was

25 repeated until the stain was removed from the gel pieces. The gel pieces were dried under
vacuum for 10 to 15 minutes. Proteins were digested overnight at 37 °C in a total volume
of 25 \iL of sequence-grade, modified trypsin (Roche Diagnostics, Indianapolis, IN) at a
final protein of 25 ng/^L in 25 mM ammonium bicarbonate. Peptides were eluted with 50%
acetonitrile and 0.5% trifluoroacetic acid. All peptide samples were concentrated, desalted,

30 and detergents removed by using CI 8 reversed-phase ZipTip™ pipette tips as described by
the manufacturer (Millipore, Bedford, MA, cat. no. ZTC18S096).

WO 03/039483 PCT/US02/36028
The resulting tryptic peptides were analyzed directly by mass spectrometry. Mass
spectrometry experiments were carried out on a PerSeptive Biosystems (Framingham, MA)
Voyager DE-STR equipped with a N 2 laser (337 nm, 3-nsec pulse width, 20-Hz repetition
rate). The mass spectra were acquired in the reflectron mode with delayed extraction.
Internal mass calibration was performed with low-mass peptide standards, and mass-
measurement accuracy was typically ±0. 1 Da. All peptide samples were diluted in a-
cyano-4-hydroxycinnamic acid, which had been prepared by dissolving 10 mg in 1 mL of
aqueous 50% acetonitrile containing 0.1% trifluoroacetic acid.

Several tryptic peptide masses from S-ADH were further sequenced by one of the following
approaches by mass spectrometry as described below.

Approach 1: Tryptic digests of the protein were derivatized with chlorosulfonylacetyl
chloride reagents as described by Keough T., Lacey M.P., YQungquist R.S. Proc. Natl
Acad. Scl USA 1 999; 96 7 1 3 1 . The sulfonated sample was acidified with trifluoroacetic
acid and cleaned up directly using CI 8 mini-columns (ZipTips™, Millipore). The
derivatized peptides were eluted into a-cyano-4-hydroxycinnamic acid (Fluka, cat. no.
28480) and plated directly onto MALDI plates. Derivatized peptides were analyzed on an
Applied Biosystems Voyager DE-STR time-of-flight mass spectrometer equipped with a N 2
laser. All mass spectra were acquired in the reflectron mode with delayed extraction.
External mass calibration was performed with low-mass peptide standards, and mass
measurement accuracy was typically + 0.2 Da. PSD fragment ion spectra were obtained
after isolation of the appropriate derivatized precursor ions using timed ion selection.
Fragment ions were refocused onto the final detector by stepping the voltage applied to the
reflectron in the following ratios: 1.0000 (precursor ion segment), 0.9000, 0.7500, 0.5625,
0.4218, 0.3164, and 0.2373 (fragment ion segments). The individual segments were
stitched together using software developed by Applied Biosystems. All precursor ion
segments were acquired at low laser power (variable attenuator = 1980) for 100 laser pulses
to avoid detector saturation. The laser power was increased (variable attenuator = 2365) for
the remaining segments of the PSD acquisitions. The PSD data were acquired at a
digitization rate of 20 MHz; therefore, all fragment ions were measured as chemically
averaged and not monoisotopic masses.

WO 03/039483 PCT/US02/36028

Approach 2 : Sequence tags were obtained from S-ADH tryptic peptides. Post source decay
(PSD) fragment ion spectra were acquired for four peptides after isolation of the appropriate
precursor ion by using timed ion selection. Fragment ions were refocused onto the final
detector by stepping the voltage applied to the reflector in the following ratios: 1.0000
5 (precursor ion segment), 0.9000, 0.7500, 0.5625, 0.4218, 0.3164, and 0.2373 (fragment
segments). The individual segments were stitched together by using software provided by
PerSeptive Biosystems. All precursor ion segments were acquired at low laser power
(variable attenuator = 1 ,450) for <256 laser pulses to avoid saturating the detector. The
laser power was increased for all of the remaining segments of the PSD acquisitions.
!0 Typically, 200 laser pulses were acquired for each fragment-ion segment. The PSD data
were acquired at a digitization rate of 20 MHz. Mass calibration was performed with
peptide standards. Metastable decompositions were measured in all PSD mass spectrometry
experiments.

15 Approach 3 : Sequence tags were obtained from S-ADH tryptic peptides by ESI MS/MS the
mass spectra were acquired on a Micromass Q-TOF2 quadrupole/time of flight MS system.

Example 3

Initial electrochemical measurement ofNADHand correlation to spectrophotometric data,
20 1 0 micron disc carbon fiber microelectrodes were purchased (from Bioanalytical Systems
("BAS"), West Lafayette, Indiana (part number MF-2007)) and pretreated using the method
of Kuhr et. al (63). The electrode surface was polished for 10 min. with 1 \im diamond
paste (Bioanalytical Systems) and sonicated in hot toluene for 2 min. To remove residual
polishing material, the microelectrode was rinsed once in methanol and once in water, then
25 sonicated twice in water for 1 min. The polished microelectrode was subsequently

pretreated electrochemically in 1 M HC1 by twice applying 10 cycles of 100 V/s from -200
mV to +1800 mV. Then the microelectrode was treated in 100 mM potassium phosphate
buffer by twice applying 10 cycles of 0 to +1200 mV at 100 mV/s. Background scans were
then obtained from phosphate buffer alone. All potentials were referenced versus a
30 Ag/AgCl reference electrode (Bioanalytical Systems). After baseline fast-scan cyclic

voltammograms (CVs) were obtained for the enzyme (1 U/mL) and NAD(P)H (2 mM), the
required volume of aqueous acetone was added (20 mM final concentration). The solution
was quickly mixed, and fast-scan CVs were obtained every 1 min. for 25 min. The buffer-

WO 03/039483 PCT/USO 2/36028

only background was subtracted from each CV with BAS 100W electrochemical software
version 2.3 (obtained from Bioanalytical Systems, West Lafayette, Indiana, hereinafter
"BAS").

5 Unless otherwise indicated, all electrochemical measurements were performed using a

Bioanalytical Systems (BAS) Model 100A or B electrochemical analyzer coupled to a BAS
PA-1 preamplifier and a Faraday cage (part number MF-2500), wherein all waveforms were
generated and currents acquired via BAS 100W electrochemical software version 23. The
data were processed using Microsoft Excel 97 SR-2 and BOMEM GRAMS/32 version 4.04,
10 Level II (Galactic Industries Corporation). The electrochemical cell was a custom-built
0.20 mL cell, constructed from Plexiglas (acrylic polymer sheet, from Atofina Corp., Paris,
France), containing a Ag/AgCl reference electrode, the pretreated carbon fiber
microelectrode, and a Pt wire auxiliary electrode.

15 To correlate spectrophotometric with electrochemical data for both enzymes, the same
reaction conditions were used for both analyses. For S-ADH from T. brockii, the 1 mL
reaction volume comprised final concentrations of 2 mM NADPH, 20 mM acetone, and 1 U
S-ADH. For S-ADH fromX autotrophicus Py2, the 1 mL reaction volume comprised final
concentrations of 2 mM NADH, 20 mM acetone, and 1 U S-ADH. For both reactions,

20 baseline A 34 o was obtained for the enzyme and NAD(P)H versus a phosphate buffer blank.
The cuvette containing the reaction solution was then removed from the spectrophotometer,
and the 0.4 mL of solution was removed from the cuvette and combined with the remaining
0.6 mL. The required volume of aqueous acetone was added to the 1.0 mL reaction. The
solution was mixed, 0.4 mL was added to the cuvette, and the cuvette replaced in the

25 spectrophotometer. The decrease in A340 was then monitored for 30 min. using a Shimadzu
UV-VIS-NIR scanning spectrophotometer (model UV-3101PC, Colombia, MD). Data were
acquired using UVPC Personal Spectroscopy Software version 3.9 (Shimadzu, Colombia,
MD) and processed using Microsoft Excel 97 SR-2. Quartz cuvettes with a 1 mm
pathlength and a 0.4 mL volume were purchased (from Fisher Scientific, Pittsburgh,

30 Pennsylvania, part number 14-385-906A).

. Electrochemical measurement of acetone-dependent NADH consumption using Meldola 's
Blue-modified carbon electrodes, A glassy carbon disk electrode modified with the

WO 03/039483 PCT/US02/36028
electrocatalyst Meldola's blue was prepared as follows. A 3-mm diameter glassy carbon
electrode (BAS part number MF-2012) was first wet-polished with a 1 \im diamond
suspension, sonicated in deionized water for one minute, and then further polished with 0.05
|im alumina polishing suspension. The freshly polished electrode was.washed thoroughly
5 by sonication in deionized water and subsequently pretreated electrochemically in 5 mL
deoxygenated 100 mM phosphate buffer (pH 7.2) by applying 20 cycles of 5 V/s from -500
mV to +300 mV, four times. After the cycling, a constant polarizing potential at -0.5 V
was applied for 60 s. The electrochemically pretreated electrode was then soaked in 0.5%
of Meldola's blue (Aldrich, Milwaukee, WI, catalog number 32,432-9) at room temperature
10 for 30 min. The electrode was rinsed with deionized water before use.

Screen-printed carbon electrodes formulated with Meldola's Blue mediator were purchased
from Gwent Electronic Materials Ltd. (Pontypool, United Kingdom). The disposable strips
were configured in the geometry described by Hart et ah and consisted of two screen-

15 printed electrodes deposited onto a polyethylene substrate. The working electrode was

graphite carbon containing the electrocatalyst Meldola's Blue (part number C70902D2 from
Gwent), and the reference/counter electrode was Ag/AgCl printed ink (part number
C61003D7 from Gwent). The working electrode area was defined by printing an additional
dielectric coating (part number D2000222D2 from Gwent). The electrode geometric area is

20 3 x 3 mm, or 9 mm 2 . The electrodes were pre-soaked in phosphate buffer for 10 minutes
before use to remove loosely bound Meldola's Blue.

The acetone-dependent consumption of NADH catalyzed by S-ADH was measured with
Meldola's blue-carbon electrodes prepared as above using chronoamperometry in a 1 mL
25 reaction volume containing 100 mM potassium phosphate buffer (pH 7.2), NADH (500
|iM), S-ADH (1 U), and varying concentrations of acetone. After a 2 min. incubation
period, the potential was stepped from open circuit to 68 mV (vs. Ag/AgCl) and the current
was recorded after 1 20 s.

30 Measurement of acetone-dependent consumption of NADH using commercial blood glucose
disposable test strips. Disposable glucose biosensor strips and reader (Precision Xtra
Advanced Diabetes Management System) are available from MediSense (a division of
Abbott Laboratories, Bedford, MA). 1 mL reaction volumes containing 25 mM potassium

WO 03/039483 PCT/US02/36028
phosphate buffer (pH 6.2), NADH (2 mM), S-ADH (20 U), and acetone (0.5, 1.0, 1.5, 2.0
mM respectively) were incubated at room temperature. After 5 min., a 20-^L aliquot was
removed from each reaction mix and applied to a disposable strip pre-inserted in the glucose
meter. The meter reading value (mg/dL of glucose equivalent) was recorded and plotted to
the amount of acetone added.

Secondary alcohol dehydrogenase coupled to H 2 0 2 formation electrochemical assay. A
disk platinum electrode (BAS part number MF-2013) was used to monitor H 2 0 2 produced
by the S-ADH coupled enzymatic reaction in response to acetone concentration. Before
measurements the electrode surface was polished using A1 2 0 3 paste for 1 rnin. and then
rinsed with deionized water, sonicated for 1 min. and rinsed with water again. The polished
Pt electrode was then pretreated electrochemically by applying 10 cycles of 100 mV/s from
+200 mV to +900 mV. All potentials were referenced versus a Ag/AgCl electrode (BAS
part number MF-2078). Assays contained potassium phosphate (100 mM, pH 7.2), purified
S-ADH (1 U/mL), NADH (20 ^M), lactate (100 mM), lactate dehydrogenase (5 U/mL),
pyruvate oxidase (4 U/mL), flavin adenine dinucleotide (0.01 mM), cocarboxylase (0.2
) mM), in a total volume of 0.5 mL. Assays were initiated by addition of acetone. After a 2
min. incubation period, the potential was stepped from open circuit to 350 mV. The
oxidative current was recorded after 120 s and plotted against acetone concentration.

Disposable electrode materials were evaluated to monitor acetone-dependent H 2 0 2
produced by the coupled enzyme reaction using the identical enzyme reagent system and
similar electrochemical technique as described above for the disk platinum electrode.
Screen-printed platinized carbon/graphite electrodes and cobalt phthalocyanine carbon
electrodes were purchased (part numbers C200051 1D1, and C4051 1D8, respectively,
Gwent Electronics Materials, Ltd.) with the same electrode geometry as described earlier
for the Meldola's Blue screen-printed carbon electrodes. Screen-printed platinized carbon
electrodes were pre-soaked in phosphate buffer for 5 min. before use. Assays were initiated
by addition of acetone and incubated for 2 min. at which time the potential was stepped
from open circuit to 350 mV. The oxidative current was recorded after 120 s. Cobalt
phthalocyanine-modified screen-printed carbon electrodes were pre-soaked in phosphate
buffer for 5 min. before use. After each addition of acetone, the reaction was allowed to
incubate for 3.5 min. Chronoamperometric measurements were made with an initial quiet

WO 03/039483 PCT/USO 2/36028

time of 5 s at 150 mV, and then the potential was stepped to 650 mV for 30 s and the
current recorded. One cobalt phthalocyanine-modified screen-printed electrode was used
for each experiment and then discarded.

5 A prototype disposable platinized carbon electrode was constructed by cutting 1/8 inch
(3.06 mm) diameter circular disks (using a manual hole puncher) of Toray carbon paper
(porous carbon paper) or cloth, loaded with 20% (w/w) platinum nanoparticles (these
platinum particles are nanonoparticles deposited on carbon; the platinum nanoparticle-
loaded paper or cloth was purchased from ETEK Division of De Nora North America,

10 Somerset, NJ, part number SLS-SPEC) and attached to a screen-printed carbon working
electrode (part number C10903D14 from Gwent Electronics Materials, Ltd.) using double-
sided carbon tape (also 1/8 inch (3.06 mm) diameter disk). In some experiments, 20 \iM of
non-ionic detergent TRITON X-100 (t-octylphenoxypolyethoxyethanol; catalog number T-
8787, from Sigma Chemical Co.) or BRIJ 30 (tetraethylene glycol monododecyl ether;

15 catalog no. P-1254, from Sigma) was applied to the ETEK material disk and allowed to dry
before use. Before measurements, the electrode was pretreated electrochemically by
applying 10 cycles of 100 mV/s from +200 mV to +900 mV twice. Assays were initiated
by addition of acetone and incubated for 2 min. Chronoamperometric measurements were
made with a quiet time of 2 s at 215 mV, and then the potential was stepped from 215 mV

20 to 350 mV vs. Ag/AgCl. The oxidative current was recorded after 30 s.

Reflectance photometry measurement of acetone-dependent H2O2 formation using glucose
disposable test strips and correlation to electrochemical data. Disposable glucose
biosensor strips and reader were purchased (OneTouch Basic read and strips from Lifescan,

25 Inc., Milpitas, California). Successive additions of 100 |iM acetone were added to a 1 mL
reaction volume containing the S-ADH coupled enzyme system (as described above) and
incubated at room temperature. Each acetone addition was allowed to react for 4 min. and
then a 20-^L aliquot was removed from the reaction mix and applied to a disposable strip
pre-inserted in the glucose meter. The meter reading value (mg/dL of glucose equivalent)

30 was recorded and plotted against the total concentration of acetone. H 2 0 2 concentration
was also monitored chronoamperometrically using a disk platinum electrode as described
above. The correlation between the electrochemical assay and the colorimetric readings
were plotted.

WO 03/039483

PCT/US02/36028

Enzyme-based electrochemical measurement of gas phase acetone. Gas phase samples (0-
] 0 ppm v/v) of acetone were prepared by injecting standard concentrations of acetone into a
calibrated airbag (10 L bag, Calibrated Instruments, Inc, Ardsley, N.Y.) filled with 7 L of
water-saturated air and 1 L of dry air, and allowed to evaporate at 37 °C (about 30 min.).
The gas samples produced from this system closely simulate human breath in terms of
temperature and moisture content The gas sampling system was calibrated (that is,
concentration of acetone gas phase and liquid phase samples) using gas chromatography
with a Hewlett Packard 5890 gas chromatograph equipped with flame ionization detection
and an on-column injector. 1 jiL aqueous samples were applied to a 1 5m long, coiled
capillary column (Nukol, 0.53mm diameter with 0.50-nm layer of liquid phase, catalog
number 25326, available from Supelco, Inc., Bellafonte, Pennsylvania). The oven
temperature was held at 40°C for 4 min., then increased at 25°C/min. to 200°C The carrier
gas flow rate was 5mL/min. of helium.

Two types of sampling techniques were used to partition acetone from the gas phase into the
liquid phase; a foam system, and a thin-aqueous layer system. For the foam system, a piece
of polyurethane foam was cut into a cylindrical shape (19 mm long and 10 mm in diameter)
so that the volume was about 1 mL. The foam was boiled in water for 20 min. and then
inserted into a 3 cc disposable plastic syringe. The syringe plunger was inserted and pushed
firmly to remove excess water and then removed. Before introducing gas phase acetone
samples, 50 jiL of water or phosphate buffer was loaded into the foam. Once the water
contacted foam, the surface tension sucked water into the foam cell and the water
distributed evenly onto foam surface. The syringe containing wetted foam was then
connected via tubing to the gas sampling system and the gas sample passed through the
foam with a flow rate 5 L/min. for 1 2 seconds either by running a diaphragm pump or by
manually pushing the airbag. This allowed the total gas sample volume to equal 1 L. After
sampling, the syringe containing foam was quickly disconnected and the plunger re-
inserted. The liquid was then squeezed out into an electrochemical cell for electrochemical
analysis or into a vial insert for gas chromatography analysis. For electrochemical
measurements, the acetone-partitioned water sample was mixed with concentrated enzyme
solution (S-ADH and coupling enzymes as discussed above) to make the desired final

WO 03/039483 PCT/US02/36028
enzyme solutions and incubated for 2 min. The acetone-dependent H2O2 formed from the
enzyme reaction was measured chronoamperometrically as described above.

For the thin aqueous layer sampling method, the gas was released from the airbag in a fine
5 stream at a flow rate of 500 mL/min. for 2 min. so that the total volume of gas was equal to
1 .0 L. In this experiment, the working electrode was inverted (electrode surface facing up),
so that a small amount of enzyme solution (50 uL) forms a relatively thin layer of liquid to
cover the electrode surface. The gas was blown perpendicular to the liquid surface. The
gas stream stirred the liquid to enhance the mass transfer of acetone from gas phase into
10 liquid phase. After the gas sample flow, the enzyme solution was allowed to react for 1
min. The acetone-dependent H2O2 formed from the enzyme reaction was measured
chronoamperometrically as described above. The current responses were plotted against the
gas-phase acetone concentration in the airbag.

15 Data analysis. Kinetic constants (K m and V mttK ) were calculated by fitting initial rate data to
the Michaelis-Menten equation and using the software KaleidaGraph Fourth Edition,
(Synergy Software, Reading, PA). .

RESULTS

Purification ofS-ADHfromX. autotrophic™ strain Py2. S-ADH enzymes are believed to
20 be involved in primary (growth on isopropanol) or intermediary steps (growth on propane)
in bacterial pathways involving aliphatic hydrocarbon catabolism. These enzyme reactions
are freely reversible where the reaction direction is dependent on the chemical equilibrium.
The reverse reaction, that is the reduction of acetone to isopropanol with concomitant
oxidation of NADH or NADPH (most dehydrogenases have preference for one coenzyme
25 over the other), arguably provides the substrate specificity and redox chemistry necessary
for an enzyme-based electrochemical sensor. For these reasons, this class of enzymes was
investigated in more detail by testing for S-ADH activity in cell extracts of isopropanol-,
propane-, and acetone-grown bacterial cultures (data not shown; growth screens and isolates
performed as described in Experimental Procedures). Of these, cell extracts of isopropanol-
30 grown X. autotrophicus strain Py2 had the highest S-ADH specific activity (1 .2 umol

acetone reduced min ^mg" 1 protein) and also preliminarily observed to have specificity for
secondary alcohols over primary alcohols. To investigate the properties of this enzyme

WO 03/039483 PCT/US02/36028
further, S-ADH was purified on the basis of its ability to reduce acetone in the presence of
NADH or oxidize isopropanol in the presence of NAD + . The enzyme was purified about
40-fold using DEAE-Sepharose and RED SEPHAROSE chromatography with an estimated
recovery of 30 to 40%. As shown in Figure 2, this two-step purification resulted in the
enrichment of a single polypeptide with an apparent molecular mass of 42 kDa on SDS-
PAGE. A polypeptide band that migrated with the same apparent molecular mass was
weakly visualized in cell extracts of isopropanol-grown Xanthobacter that was not readily
visible at this same position in cell extracts of glucose-grown Xanthobacter (Figure 2).
Consistent with SDS-PAGE analysis, S-ADH activity in glucose-grown cell extracts was
only 3% of the activity observed in isopropanol cell extracts, suggesting that S-ADH is
induced by the presence of isopropanol. Mass spectrometry provided a more accurate
molecular mass estimate of 37. 1 kDa for purified S-ADH, which is consistent with observed
apparent molecular mass determined by SDS-PAGE. The pi of S-ADH was determined to
be 7.4 by isofocusing electrophoresis. The N-terminal sequence of the S-ADH polypeptide
was determined to be MKGLVYRGPGKKALE (SEQ ID NO:7) by Edman degradation.
Additional peptide amino acid sequences were determined by digesting the polypeptide with
trypsin to produce "tryptic fragment" and employing matrix-assisted laser desorption
ionization-mass spectrometry using post-source decay sequencing (PSD-MALDI) to
determine their amino acid sequences.

The amino acid sequences of six peptides were obtained by derivatizing the fragments
followed by sequence analysis using PSD-MALDI (Table 2).
Table 2. Sequences of derivatized tryptic fragments of Xanthobacter Py2 S-ADH.

Observed MH+ (m/z)

Amino Acid Sequence

1431.33

PVAVDHGP(FS)PHK (SEQ ID NO:8)

1111.1

GG(L/I)GVYHQ (SEQ ID NO:9)

1047.2

A(L/I)EEVPHPR (SEQ ID NO: 10)

1047.1

HPSGDTR (SEQ ID NO:l 1)

946.27

GLVYRGPGK (SEQ ID NO: 12)

756.17

HQ(I/L)ASSR (SEQ ID NO: 13)

The sequence of the peptide with the observed mass-to-charge ratio of 946.27 is identical to
a nonamer present within the N-terminal sequenced derived by Edman degradation. Using
PSD MALDI analysis, an additional four amino acid sequences were detected (Table 3).

WO 03/039483 PCT/US02/36028
Table 3. Sequences of Xanthobacter Py2 S-ADH fragments, by PSD-MALDI.

Observed MH+ (m/z)

Amino Acid Sequence

1151

LDNVPE (SEQ ID NO: 14)

1331

FDQRQP (SEQ ID NO: 15)

2288

GAGRIIAV (SEQ ID NO: 1 6)

2422

QVEPLMS (SEQ ID NO: 17)

The amino acid sequences of S-ADH tryptic peptide fragments were also obtained by ESI
MS/MS (Electrospray Ionization, tandem Mass Spectrometry/Mass Spectrometry). The
resulting MS/MS analysis was consistent for the following amino acid sequences (Table 4).
Table 4. Sequences of Xanthobacter Py2 S-ADH fragments, by ESI MS/MS.

Observed MH+ (m/z)

Amino Acid Sequence

1760.0

FFADIIEAA (SEQ ID NO : 1 8)

1330.8

DTVTTH (SEQ ID NO: 19)

Thus, the Xanthobacter Py2 S-ADH was characterized as a protein having NAD + -dependent
secondary alcohol dehydrogenase activity, having the ability to reduce acetone to

10 isopropanol, and having specific activity for ketones and secondary alcohols; having, for the
oxidation of isopropanol to acetone, (1) a pH optimum of approximately 7.8, and (2) an
average specific activity ratio for secondary-to-primary alcohols of at least 50:1 when tested
at pH 7.8 under equivalent conditions individually with C3-C5 straight chain secondary
alcohols and with C2-C5 straight chain primary alcohols; having, for the reduction of

15 acetone to isopropanol, (3) a pH optimum of approximately 6.2, (4) an apparent K m of

approximately 144 ± 18 |iM, (5) an apparent K max of approximately 43.4 ± 1.2 \xmo\ acetone
reduced-min^-mg" 1 protein, (6) an apparent £ cat of approximately 30.4 sec" 1 , (7) an apparent
kcai/K m of approximately 2.1 x 10 s , and (8) a K m for NADH of approximately 5.1 ± 0.4 ^M;
and comprising at least one polypeptide molecule that has (a) a molecular mass of

20 approximately 37.1 kDa as determined by mass spectrometry, (b) a pi of approximately 7.4
as determined by isofocusing electrophoresis, and (c) a tetradecameric N-terminal amino
acid sequence of SEQ ID NO:7, and that is capable of being degraded to form fragments
having the amino acid sequences of SEQ ID NO:8 to SEQ ID NO: 19.

25 In addition, a terminally-truncated version of this protein was also co-purified. This

terminally-truncated version was missing a terminal decapeptide and yet functioned as well
as the intact protein (data not shown). The term "enzyme" as used herein, means
"catalytically functional biomolecule," which includes both whole native (or native-size)

WO 03/039483 PCT/US02/36028

molecules, as well as terminally truncated versions retaining function, that is having up to
about 10 amino acids deleted from at least one terminus. In specific reference to the
Xanthobacter Py2 S-ADH, the term "enzyme" therefore includes both the whole native
molecule and the terminally truncated versions.

Substrate specificity and inhibition of S-ADH from X. autotrophicus strain Py2. In acetone
reduction assays, NADH but not NADPH could serve as the electron donor (data not
shown). The pH optimum for reducing acetone to isopropanol was 6.2 while the optimum
for the reverse reaction was 7.8 (data not shown). Kinetic constants were calculated from
data collected from assays in which the NADH or acetone concentration was held at a fixed
saturating concentration, while the acetone or NADH concentration was varied in individual
assays. The apparent K m9 K max , £ cat , and hJK m values were calculated to be 144 ± 18 \iM,
43 .4 ± 1.2 jimol acetone reduced-min ^mg" 1 protein, 30.4 sec' 1 , and 2.1 x 10 5 , respectively.
The K m for NADH was calculated to be 5.1 ± 0.4 p,M. Of several compounds evaluated as
alternative substrates, S-ADH exhibited specificity for ketones and secondary alcohols
(Figures 3 and 4). In Figure 4, "NA" indicates "no activity," that is that no activity toward
the substrate was detected. As shown, primary alcohols exhibited no activity or very low
activity in comparison. Among the ketones and secondary alcohols, the highest specific
activities were observed with 2-pentanone and 2-pentanol for the reductive and oxidative
reactions, respectively.

As a result, the only tested primary alcohol toward which the enzyme exhibited activity was
1-butanol and the specific activity measured (at pH 7.8) was only 1.7 ± 0.2 jimol butanol
oxidizedmin '-mg' 1 protein. No activity toward ethanol, 1-propanol, or 1-pentanol was
detected. The average specific activity toward C2-C5 straight chain primary alcohols was
thus, at most, about 0.475 ^mol alcohol oxidizedmin^-mg" 1 protein. Activity toward
secondary alcohols was tested for C3-C5 straight chain secondary alcohols. The enzyme
exhibited activity toward all three, and the specific activity values measured (at pH 7.8)
were 24 ± 1 .8 )xmo\ 2-propanol oxidized-min^-mg 1 protein, 21 ± 3.6 nmol 2-butanol
oxidized-min^-mg" 1 protein, and 37 ± 1.1 jimol 2-pentanol oxidized min" ^mg* 1 protein. The
average specific activity toward C3-C5 straight chain secondary alcohols was thus, at least,
about 25 jimol alcohol oxidized-min ^mg" 1 protein. Therefore, for this S-ADH, the
minimum average specific activity ratio for secondary-to-primary alcohols, when tested at

WO 03/039483 PCT/US02/36028

pH 7.8 under equivalent conditions individually with C3-C5 straight chain secondary
alcohols and with C2-C5 straight chain primary alcohols, may be given by 25/0.475 which
equals 52.6; thus, this S-ADH enzyme, tested under these conditions, has an average
specific activity ratio for secondary-to-primary alcohols of at least about 50:1.

Using saturating concentrations of acetone, no inhibition was observed for S-ADH activity
in the presence of methanol, ethanol, and 1-propanol at concentrations about 13 -fold over
the K m value for acetone (data not shown). S-ADH activity was tested in the presence of
higher concentrations of ethanol since this compound is a key potential inhibitor for
10 diagnostic breath acetone analysis. S-ADH was 82-85% active in the presence of 41 mM
ethanol which is equivalent to 500 ppm (v/v) gas phase ethanol and is the upper limit
concentration of ethanol found in human breath.

The acetone detection limit of S-ADH was investigated using spectrophotometry in order to
15 assess this enzyme's ability to detect the low levels of acetone present in human breath.

Using a high loading of enzyme (4 U), the NADH consumption response time was less than
20 s and linear (correlation coefficient = 0.99997) for acetone concentrations ranging from
0.058 to 5.8 ppm (w/v). The lower end of the acetone detection range (0.058 to 0.29ppm
(w/v)) is within the detection parameters required for diagnostic breath acetone analysis
20 (lower breath acetone level is 0.3 to 0.8 ppm (w/v) for non-dieting individuals).

Furthermore, this detection method is limited in sensitivity, therefore other detection
methods (for example, electrochemical) may provide increased sensitivity for detecting
lower concentrations of acetone using the S-ADH reaction.

25 Stability of S-ADH from X. autotrophics strain Py2. Stability of the enzymes that are

incorporated into biosensors is one of the main parameters that determine their commercial
viability. Maintaining enzyme activity can influence storage costs, performance, and
manufacturing processes. Therefore it was necessary to find conditions for stabilizing S-
ADH activity for long periods at room temperature, preferably in a dry form. Enzyme

30 stability can be enhanced by the presence of compatible solutes that maintain the ionic and
hydrophilic environment surrounding the enzyme during lyophilization and storage. As can
be seen in Figure 5, purified S-ADH was lyophilized and found to retain approximately 60
to 70% activity relative to a non-lyophilized control when re-suspended and assayed within

WO 03/039483 PCT/US02/36028

one day of lyophilization. The presence of the disaccharide trehalose was of considerable
benefit in maintaining activity, where samples held at room temperature for 65 and 213 days
retained approximately 100% and 70% activity, respectively S-ADH samples containing no
additives were completely inactive after 65 days at room temperature. In a separate
5 experiment, S-ADH was included into a mixture of compounds that closely resembles a
formulation recently described for fabrication of a thick-film, screen-printed enzyme
electrode. This formulation included the additives hydroxyethyl cellulose (2% w/v),
DEAE-Dextran (10 mg/mL), and lactitol (10 mg/mL) and the mixture was allowed to air
dry as opposed to being lyophilized. Using this mixture, S-ADH retained 100% of its
10 activity after being stored at room temperature for 23 days (data not shown).

Biochemical comparison of S-ADH from X autotrophicus strain Py2 to other S-ADH
enzymes. S-ADH fxomX. autotrohpicus strain Py2 represents the first S-ADH enzyme
purified from a C02-dependent acetone-utilizing organism. This enzyme is similar in

1 5 several respects to previously described secondary alcohol dehydrogenases in terms of its
molecular mass and substrate specificity. The NAD-dependent secondary alcohol
dehydrogenases from Pseudomonas sp. 6307 (ATCC 21439), Mycobacterium vaccae strain
JOB-5 (ATCC 29678), Candida boidinii (ATCC 32195), Rhodococcus rhodochrous
PNKbl (Ashraf & Murrell (1990)), and the NADP-dependent S-ADH from

20 Thermoanerobium brockii all have subunit molecular masses ranging from 37 kDa to 48
kDa. S-ADH purified from M vaccae strain JOB-5 is a 37 kDa molecular weight (subunit)
and is induced by growth on propane. In contrast to the Xanthobacter S-ADH, the M.
vaccae enzyme appears to have some specificity for primary alcohols in addition to
secondary alcohols, although primary alcohols have significantly higher K m values (for

25 example, K m = 0.05 mM vs. K m = 8.1 mM for 2-propanol and 1-propanol, respectively).
Comparatively, the K m for acetone reduction was calculated to be 0.3 mM. Another
difference is the pH optimum for the reduction and oxidation reactions. M. vaccae JOB-5
has a pH optimum of 10-10.5 for oxidizing 2-propanol and 7.5-8.5 for reducing acetone.
Another secondary alcohol dehydrogenase was isolated from the propane-utilizing

30 bacterium R. rhodochrous PNKbl (Aft = 42 kDa), although this enzyme exhibited nearly
equal specificity for either secondary or primary alcohols (for example K m = 18 mM vs. K m
= 12 mM for 1-propanol and 2-propanol, respectively). In contrast to the S-ADH enzymes
found in propane utilizers, S-ADH from the methylotroph Pseudomonas species 6307

WO 03/039483 PCT/US02/36028

(AATCC 21439) (Mr = 48 kDa) was demonstrated to be highly specific for secondary
alcohols and exhibited no activity with primary alcohols. It was also shown that this
enzyme was largely not inhibited in the presence of high concentrations of ethanol (0% and
30% inhibition in the presence of 10 mM and 100 mM ethanol, respectively). S-ADH from
5 Brockii (Mr = 40 kDa) exhibits very similar substrate specificity to S-ADH from

Xanthobacter with the exception that it prefers NADPH rather than NADH. Of note is that
S-ADH from T. brockii is one of the few S-ADH enzymes that has been cloned and
sequenced (Genbank accession number A32973). Other available S-ADH enzymes (S-
ADHs) include S-ADH from Candida parapsilos, as described in US 5,763,236; the

10 sequence available in Genbank, under Accession No. ABO 10636; S-ADH from

Thermoanaerobacter ethanolicus 39E (ATCC 33223) (Genbank Accession No. U49975);
and S-ADH from Clostridwn beijerinckii (Genbank accession number M84723).
Comparatively, five of the first 1 5 amino acid residues of the N-terminal amino acid
sequence (MKGFAMLSIGKVGWI) of S-ADHs from T. brockii and T. ethanolicus (both

15 enzymes have identical N-terminal sequences) match the N-terminal sequence of S-ADH
from* autotrophics st. Py2 (MKGLVYRGPGKKALE).

It is apparent from the literature that there may be many S-ADH enzymes that are specific
for acetone. Such S-ADH enzymes could be incorporated into a biosensor. In addition to

20 these, culture collection strains and several new strains that were isolated from soil by

enrichment on acetone, isopropanol, or propane as growth substrates were found to exhibit
S-ADH activity (data not shown). In order to compare and evaluate relative specificities
among these S-ADH activities, purified S-ADH from Xanthobacter and partially purified
preparations of, or cell extracts containing, S-ADHs from isopropanol-grown bacterial

25 strains (TDCC IP-1 (and two additional strains: data not shown)), propane-grown M vaccae
JOB-5, and T. brockii S-ADH were assayed with various ketone and alcohol substrates. As
shown in Table 5, S-ADH enzymes from X autotrophics Py2, strain IP-1 , M. vaccae JOB-
5, and T. brockii all demonstrated negligible or substantially lower activity with primary
alcohol substrates. Although the S-ADH reaction equilibrium for acetone monitoring will

30 be shifted for acetone reduction (that is NAD(P)H in excess), low specificity for primary
alcohols, mainly ethanol, is preferred to alleviate possible inhibitory effects. All of the
enzymes were active using longer-chain ketones (that is 2-butanone, 2-pentanone, and 3-
pentanone) however this should be of little consequence since these compounds are not

WO 03/039483 PCT/US02/36028

normally found on human breath. Interestingly, S-ADH from T. brockii was more active
with shorter chain ketones, which contrasts the activity of S-ADH from Xanthobacter which
was more active with longer chain ketones. It should be noted that the T. brockii S-ADH is
a commercially available S-ADH (Sigma). The application of this enzyme in a biosensor
according to the invention will be discussed in more detail below. Thus, the X.
autotrophics S-ADH enzyme isolated as described above represents a unique enzyme that
enables the relatively selective detection of longer chain ketones in mammalian samples via
the coupled enzyme system of the invention.

Table 5. Substrate Specificities of Bacterial Secondary Alcohol Dehydrogenases.

Substrate

Relative S-ADH Activity (%) ol

" Bacteria] Strains

X. Py2

IP-1

JOB 5

T. brockii

C. boidinii*

Acetone 8

100

2-Butanone

170

192

213

175

2-Pentanone

198

185

3-Pentanone

180

Acetone/Ethanol b

100

2-Propanor

100

Ethanol

NA d

1-Propanol

1-Butanol

2-Butanol

147

130

121

100

1-Pentanol

2-PentanoI

157

Ketone reduction rates calculated in duplicate at pH 6.2 with 2.5 mM substrate and 0.2
mM NADH (NADPH for T. brockii assays).

0 Acetone reduction rates calculated in duplicate at pH 6.2 in the presence of an equivalent
amount of ethanol (2.5 mM).

c Alcohol oxidation rates calculated in duplicated at pH 7.6 with 2.5 mM substrate and 0.2
mM NAD + (NADP + for T. brockii assays).

^ NA, no activity detected.

e Ketone reduction rates and alcohol oxidation rates calculated at pH 7.6 with 5 mM
substrate and 0.2 mM NADH or NAD + .

Monitoring of the S-ADH reaction electrochemically. Electrochemical transduction of
enzyme-substrate interactions provides a general analytical means to detect the respective
substrate. As previously described, dehydrogenase-based enzyme reactions typically
require the pyridine nucleotide cofactors NADH or NADPH in stoichiometric amounts for
catalysis. A number of biosensors have been described in the art which rely on
electrochemical detection of these coenzymes. As shown in Figure 6, operating in a manner

WO 03/039483 PCT/US02/36028

similar to other biosensors, electrochemical detection of NAD(P)H consumption would
allow the conjugation of an electrode and an S-ADH catalyzed reaction, providing a means
to quantify acetone in a substrate-specific fashion. In order to validate the scheme shown in
Figure 6, fast-scan cyclic voltammetry (sweeping rate >100 V/s) was performed using a
5 carbon microelectrode to measure the acetone-dependent consumption of NADPH catalyzed
by S-ADH from T. brockii. Electrochemical detection of acetone-dependent oxidation of
NADPH catalyzed by T. brockii S-ADH was measured by performing continuous
voltammetric scans of the enzyme reaction mix. These scans demonstrated changes in the
anodic peak area over time, which were dependent upon the addition of acetone. The

10 anodic current decreased versus time indicating that NADPH was being consumed
(oxidized) as S-ADH catalyzed the reduction of acetone to isopropanol. Scans were
performed from about 1200 mV to 0 mV and anodic current was measured; data for 19
scans were obtained at one-minute intervals. The initial current (at about 1200 mV) for
each scan varied regularly, with about -6.5 nA being measured at 1 minute and about -0.2

15 nA at 19 minutes. For each scan, a significant current ceased (and about 0 nA was
measured thereafter), by about 600 mV. In order to further confirm that NADPH
consumption was being accurately measured, the reaction was monitored both
electrochemically and spectrophotometrically under identical reaction conditions. As
shown in Figure 7, response data obtained using both detection methods were in close

20 agreement to each other confirming that the acetone-specific, NADPH-dependent S-ADH
reaction can be monitored electrochemically with accuracy. The voltammetric scanning
procedure used to test Z brockii S-ADH was repeated using S-ADH from A! autotrophicus
Py2 to ensure that an NADH-dependent S-ADH reaction could be monitored
electrochemically. As with the NADPH-dependent enzyme, continuous scans of the

25 reaction mix containing S-ADH from X. autotrophics Py2 yielded an acetone-dependent
decrease in anodic current over time corresponding to the enzyme-catalyzed oxidation of
NADH, as shown herein in Figure 8.

The electrochemical detection of the NAD(P)H-dependent S-ADH reaction as described
30 above provides the necessary fundamental analytical means to implement certain design
criteria to the enzyme-electrode structure. In general, dehydrogenase-based biosensors
described in the art are comprised of a conductive electrode coated with a mixture, or layers,
of a catalytically enzyme(s) and a mediator compound. When the coated electrode is

WO 03/039483 PCT/US02/36028
contacted with a sample containing the substrate for which the enzyme(s) exerts a catalytic
effect, the mediator compound transfers charge to the electrode to give a readout signal
against a reference electrode. The active electrode is preferably formed of conductive
carbon that can formulated Into screen-printable ink or other formulations/compositions that
5 enable low-cost manufacturing methods and end-use disposability. Mediator compounds
described in the art specific for NAD(P)H include viologen derivatives, quinone derivatives,
phenazine, osmium phendione, thionine, alizarin green, and Meldola's Blue (Katakis &
Dominguez, Mikrochim. Acta 126:1 1-32 (1997)). Mediators, as mentioned previously,
improve the overall sensitivity of the electrode by enhancing NAD(P)H electrooxidation
10 kinetic rates at reduced potentials. The feasibility of using the mediator Meldola's Blue
(MB) for detecting the S-ADH reaction electrochemically (as shown in Figure 9) was
investigated.

Chronoamperometry was performed using a glassy carbon electrode with absorbed
15 Meldola's Blue to measure NADH concentration. As shown in Figure 10, the current

response using the modified electrode is linear with respect to the concentration of NADH
present. This linearity was also tested and determined up to 1 mM NADH (data not shown).
Since the consumption of NADH is proportional to the concentration of acetone (that is the
reaction catalyzed by the S-ADH), these results demonstrate that Meldola's Blue modified
20 carbon electrode provides a mediated electrode system for quantifying of acetone. As a
means to investigate this further and to permit the construct a practical, disposable acetone
biosensor, chronoamperometry experiments were performed using screen-printed carbon
electrodes impregnated with the mediator Meldola's Blue (hereinafter "MB-SPCE").
Shown in Figure 1 1 is the response data of the acetone-dependent S-ADH reaction obtained
25 using MB-SPCE disposable electrode strips. The correlation coefficient (R 2 ) of the acetone
response was 0.965 and the electrode-to-electrode reproducibility was approximately 6%.
These data confirm that the acetone-specific enzyme system can be monitored
electrochemically using a practical electrode system.

30 A practical device for acetone measurement was devised in which acetone-dependent
NADH consumption was monitored. This device employed a commercially available,
disposable, glucose biosensor system (a blood glucose monitor known to register NADH
concentration, and blood glucose test strips produced for use therewith, both manufactured

WO 03/039483 PCT/US02/36028

by MediSense, Inc., Abbott Laboratories, Bedford, MA) to measure NADH consumption
catalyzed by S-ADH in response to acetone. Shown in Figure 12 are the results,
demonstrating acetone-dependent NADH consumption measured by the portable monitor
and disposable test strips. This demonstrates the feasibility of constructing a practical
5 device for acetone measurement that uses mass-produced electrodes and monitors to register
and/or record currents for an acetone-specific enzyme system.

In contrast to electrochemically monitoring the S-ADH reaction by following NAD(P)H
consumption, it is also possible to monitor NAD(P) + formation electrochemically by

10 measuring an increase in the cathodic current density at a voltage that would be directly
proportional to NAD(P) + concentration. The disadvantage of this detection scheme is that
oxygen often gives a strong interference response at the negative voltage potentials required
for electrochemical reduction of NAD(P) + . In addition, electroreducing NAD(P) + forms
radicals that dimerize, resulting in fouling of the enzyme electrode. For these reasons,

15 electrochemical experiments involving electrochemical regeneration of NAD(P) + are often
performed anodically with electron transfer mediators that shift voltage potentials away
from interfering peaks. However, these options in the context of an acetone biosensor may
not be practical without further modification, since oxygen is always present in human
breath.

One alternative to monitoring NAD(P) + formation is to indirectly couple other enzyme
activities to the S-ADH enzyme reaction. As shown in Figure 13, NAD(P) + formation can
be coupled to lactate dehydrogenase catalysis to form pyruvate. Pyruvate is then oxidized,
by pyruvate oxidase, to produce acetylphosphate and CO2, with concomitant formation of

25 H2O2. H2O2 is then electrochemically oxidized and detected at the electrode by any of the
methods well-known in the art. In this scheme, the acetone-dependent formation of
NAD(P) + (catalyzed by S-ADH) is indirectly coupled to H2O2 production. In order to
investigate the coupling between these enzyme reactions, this reaction mix (containing
coupling enzymes and reagents) was prepared with NADH-dependent S-ADH from X.

30 autotrophics Py2 in the presence of horseradish peroxidase (HRP). HRP catalyzes the
reduction of H2O2, and the oxidation of electron donors (chromogenic dye reagents),
thereby permitting the reaction to be monitored spectrophotometrically. Figure 14
illustrates the results of the coupling reactions, where an increase in absorbance was

WO 03/039483 PCT/US02/36028

observed upon addition of acetone that corresponded to H 2 0 2 formation. The lag time
required for the coupled reactions to reach a steady state was further shortened to less than
20 s by further optimization of the reaction conditions (data not shown). The coupling
reaction was modified for NADPH-dependent S-ADH from T. brockii by replacing lactate
dehydrogenase with malic enzyme. Malic enzyme is NADP + -dependent and catalyzes the
oxidation and decarboxylation of malate to form pyruvate. Pyruvate, as described above, is
further oxidized by pyruvate oxidase to form H 2 0 2 . In this case, NADP + formation is
indirectly coupled to H 2 0 2 production. Using this coupled enzyme system, a background
increase in absorbance was observed that was not acetone-dependent (data not shown).
However, approximately a 5 -fold increase in the rate of H 2 0 2 formation over the
background rate was observed after addition of acetone. (The small, background rate of
H 2 0 2 formation (acetone independent) may be due to NADPH oxidase (diaphorase) activity
catalyzed by malic enzyme.) The pyridine-nucleotide dependency of lactate dehydrogenase
and malic enzyme in the respective, coupled S-ADH reactions (that is S-ADH from X.
autotrophic™ is N ADH-dependent and is coupled to NADH-dependent lactate
dehydrogenase; and S-ADH from T. brockii is NADPH-dependent and is coupled to
NADPH-dependent malic enzyme) may be interchanged by using site-directed mutagenesis
to alter the specificity of these coupling enzymes for NADH or NADPH (Holmberg et al.,
Protein Eng. 12(10):851-6 (1999)). Alternatively, naturally occurring homologs of lactate
dehydrogenase or malic enzyme may be used that have relaxed specificity for NADH and
NADPH (B.I. Lee et al., J Mol. Biol. 307(5):1351-62 (2001)).

Besides the two, coupled S-ADH enzyme systems described above, other coupled enzyme
systems can also be usefully employed in an acetone-specific enzyme system according to
the present invention. In the scheme shown in Figure 15, S-ADH catalyzes acetone
reduction, forming NAD(P) + , which is then reduced back to NAD(P)H by the appropriate
pyridine nucleotide dependent dehydrogenase (NADH- or NADPH-specific, depending on
the cofactor requirement of the S-ADH). The oxidized product of the dehydrogenase
reaction is then the substrate for an oxidase, which generates H 2 0 2 by oxidizing the
molecule further. Other enzymes that can be employed in this coupling scheme include, but
are not limited to: alanine dehydrogenase (EC 1 .4.1 . 1), which catalyzes the NAD + -
dependent formation of pyruvate, which is then oxidized by pyruvate oxidase; saccharopine
dehydrogenase (EC 1.5.1.7), which catalyzes the NAD + -dependent formation of lysine,

WO 03/039483 PCT/US02/36028

which is then further oxidized by lysine oxidase (EC 1 .4.3.14) to form H2O2; malic
dehydrogenase (EC 1.1.1.37), which catalyzes the NAD + -dependent formation of
oxalacetate, which is decarboxylated by oxalacetate decarboxylase (EC 4.1.1.3) to pyruvate,
pyruvate then being further oxidized by pyruvate oxidase to form H2O2; and glycerol

5 dehydrogenase (EC 1.1.1 .6), which catalyzes the N AD + -dependent formation of

dihydroxyacetone, which is then further oxidized by galactose oxidase (EC 1 .1 .3.9) to
methylglyoxal. Of these, alanine dehydrogenase was tested in an enzyme coupling scheme
to verify that such alternative coupling schemes would function in tandem with S-ADH,
wherein alanine dehydrogenase in used in place of lactate dehydrogenase. Using this

10 alternative enzyme scheme, an increase in absorbance was observed upon addition of
acetone that corresponded to H2O2 formation (data not shown).

'Non-electrochemical monitoring of the S-ADH reaction coupled to H2O2 formation.
It may also be possible to construct a practical device for non-electrochemical acetone

15 measurement. To investigate whether a practical, non-electrochemical device could be

prepared for measuring acetone by means of an acetone-specific, coupled enzyme system, a
device was constructed in order to measure acetone-dependent H2O2 formation. The device
combined a blood glucose monitor and test strips manufactured for use therewith (both
manufactured by Lifescan, Inc., Milpitas, California) with an H2C>2-producing system in

20 which S-ADH catalysis of acetone was coupled to lactate dehydrogenase and pyruvate

oxidase. In the device, the enzyme-catalyzed formation of H2O2 was coupled to peroxidase
and dye reagents, which react to produce a colored reaction product when contacted with
H2O2. The device then quantified the colored product by means of a reflectance photometry
reader. Shown in Figure 16 are the results, demonstrating acetone-dependent H2O2

25 formation measured by the portable monitor and disposable test strips. The monitor reading
was the same whether acetone (♦) (linear regression of data gave y = 0.0457x + 1 .9048) or
an equivalent amount of H2O2 (H) (linear regression of data gave y = 0.044x + 1 .3333) was
added, indicating that the enzyme system did not interfere with the device reading and that
the enzyme system was able to accurately catalyze the conversion of acetone to H2O2. This

30 demonstrates the feasibility of constructing a practical non-electrochemical device for
acetone measurement that uses mass-produced electrodes and monitors to quantitatively
register and/or record reaction product concentrations for an acetone-specific enzyme
system.

WO 03/039483

PCT/US02/36028

Electrochemical monitoring of the S-ADH reaction coupled to H 2 0 2 formation.
Electrochemical detection of H 2 0 2 has been well-studied for several years and has been
applied in several commercial applications. Thus the enzyme system as described above
(that is, S-ADH reaction coupled to H 2 0 2 formation) represents a novel means to
enzymatically monitor acetone using established electrochemical methodology. To this
end, a disk platinum electrode was used to monitor the acetone-dependent formation of
H 2 0 2 catalyzed by the S-ADH coupled to lactate dehydrogenase and pyruvate oxidase,
herein termed the "3-enzyme system", by using a chronoamperometric electrochemical
method. Figure 1 7 shows the acetone-dependent current response catalyzed by the 3-
enzyme system. The currents con-elate to acetone concentrations linearly with a correlation
coefficient of 0.997 in the acetone concentration range of 0 to 200 ^iM. Eight individual
sets of measurements were made, and the relative standard deviation (%CV, coefficient of
variation) calculated to be less than 4% in the acetone concentration range of 10 to 200 jiM.
The method of detection limit (MDL), defined as 99% of the confidence limit, is 3.3 fiM
which is sufficiently sensitive enough for measuring the equivalent amount of gas-phase
acetone in human breath. The experiment was repeated for higher acetone concentrations
(up to 500 ^M) with retention of linearity (correlation coefficient R 2 = 0.992) and
reproducibility (CV = 5% over entire concentration range; 8 measurements at each
concentration) (data not shown).

The electrochemical assay results of the 3-enzyme system were directly compared with
current responses to standard concentrations of H 2 0 2 to determine whether the 3-enzyme
system accurately converts acetone to H 2 0 2 . As shown in Figure 18, the current response
due to acetone addition (♦) or H 2 0 2 addition (H) were nearly identical, with the linear
regressions for acetone and H 2 0 2 data gave near, identical equations of y = 1.1213x +
9.8416 and y = 1.1 161x + 1 3.442, respectively. This indicates that the reaction is completed
and that acetone is stoichiometrically converted to H 2 0 2 by the 3-enzyme system.

Assays were performed as above, but in the presence of possible interference compounds.
The current responses were measured for Acetone alone, as well as for Acetone plus 1 0 mM
ethanol, Acetone plus 50 jiM isopropanol, Acetone plus 50 \M isobutanol, Acetone plus 50
\iM butanone, Acetone plus 50 ^iM pentanone (data not shown). The results demonstrate

WO 03/039483 PCT/US02/36028

that ethanol, isopropanol and isobutanol do not interfere the acetone measurement. Notably,
ethanol, which was present in 1000- fold excess (10 mM) of acetone, showed no inhibitory
effect. Ethanol is a primary volatile metabolite present in human breath. The current
responses for acetone in the presence of other ketones were increased according to the
5 concentration of other ketone added, since the S-ADH does not discriminate among these
ketone substrates (for example, a current response of 50 nM butanone is equivalent to 50
|iM acetone) in the electrochemical assay. However, these other ketones are not found in
human breath at significant levels therefore do not pose interference for the breath biosensor
application.

10..

As mentioned earlier for the electrochemical detection of acetone-dependent NADH
consumption, the 3-enzyme system as described above provides the necessary fundamental
analytical means to implement certain commercial design criteria. The active electrode is
preferably formed of conductive carbon that can be formulated into screen-printable ink
15 (that is, to form a strip electrode) or other formulations/compositions that enable low-cost
manufacturing methods and end-use disposability. Toward these end-goals, a few low-cost
electrode materials were evaluated with the 3-enzyme system as a means to demonstrate the
commercial utility of this enzyme system for detecting acetone.

20 Figure 19 shows current response vs. concentration acetone using a screen-printed

platinized carbon electrode employed with the 3-enzyme system. The current responses
correlated linearly to concentration of acetone present. The error bars were calculated based
on eight measurements made using eight electrodes (single use). The reproducibility was
dominated by electrode to electrode variation mainly due to the degree of wetting of the

25 electrode surface. It was found that pre-wetting the electrode for 5 min improved the

reproducibility significantly. Another material evaluated very similar to the screen-printed
.platinized carbon electrodes was a conductive carbon cloth or paper to which was bound
highly conductive graphite particle loaded with platinum nanoparticles (10 to 20% (w/w)
loading). The cloth or paper was hole-punched and attached to a screen-printed carbon

30 electrode and used in conjunction with the 3-enzyme system for detecting acetone. Figure
20 shows the electrode current response to H2O2 generated by acetone conversion by the 3-
enzyme system (regression of data gives y = 6. 1 601 x + 1 8.889). To aid in wetting the
surface of this electrode, some electrodes were pre-treated with surfactants before use to

WO 03/039483 PCT/US02/36028

reduce the hydrophobic nature of the graphite-Pt surface. It was determined this treatment
aided in the wetting of the electrode surface while not pacifying the sensitivity of the
electrode (data not shown). A third electrode evaluated using the 3-enzyme system was
screen-printed carbon containing the mediator cobalt phthalocyanine. Figure 21 presents
the current response curve to acetone concentration using this electrode. An offset is shown
due to the electrochemical response to NADH present in the 3-enzyme system. This effect
was further investigated where current responses were recorded at the same applied voltage
in the presence of different fixed concentrations of NADH (data not shown). Here it was
determined that the cobalt phthalocyanine modified screen-printed carbon electrode
electrodes were electroactive to NADH (at the potential for H 2 0 2 measurement) and
therefore the presence of high concentrations of NADH would interfere with acetone
measurement. However, under the reagent conditions of the 3-enzyme system, the low
concentration of NADH present offsets the current response in a fixed manner (constant) in
which this can subtracted from the current response due to acetone (H2O2) without
compromising the calibration accuracy. The results of this experiment demonstrate that
platinum or platinized-carbon electrodes are not the only electrode materials that will
function in detecting acetone-dependent H 2 0 2 formation as catalyzed by the 3-enzyme
system. Other screen-printed carbon electrodes modified with mediators for the purpose of
detecting H 2 0 2 include ferrocene and its derivatives, quinone and its derivatives, osmium
bipyridine conjugated to poly(vinylpyridine), potassium hexacyanoferrate, nickelocene,
methylene blue, methylene green, and phenazine methosulfate.

Enzyme-based electrochemical measurement of gas-phase acetone. As mentioned
previously, an important feature for the enzyme-based electrochemical sensor would be its
ability to quantify gas-phase acetone concentrations. Since the enzyme electrode is
responsive in the liquid phase alone, two sampling systems were investigated that would
allow acetone gas to partition into the aqueous phase, thus allowing the sample to be
directly or indirectly introduced to the enzyme-electrode system where a current response it
then generated (proportional to the concentration of acetone present). The principle of both
sampling systems (discussed in more detail below) follow Henry's Law constant for gas-
liquid partitioning:

kh = c a • pg" 1

WO 03/039483 PCT/US02/36028

Where kh is Henry's Law constant for acetone (24 M-atm \ as reported by NIST (The
National Institute of Standards and Technology)), c a is the concentration of acetone in the
liquid phase (M), and p g is the partial pressure of acetone in the gas phase (atm). As a
result, when gas-phase acetone samples are in contact with a liquid phase, the gas-liquid
5 acetone concentration reaches equilibrium as expressed by Henry's Law. The concentration
range of gas-phase acetone in human breath that needs to be quantified by the biosensor is
0.5-10 ppm (v/v). Therefore, under equilibrium conditions, 0.5-10 ppm (v/v) gas-phase
acetone is equal to about 10 to 200 uM acetone in the aqueous phase. The basis of this
correlation was verified with gas chromatography by passing standard concentrations of

10 gas-phase acetone through a piece of polyurethane foam that was pre- wetted with a fixed
volume of aqueous buffer. Figure 22 is a diagram of the apparatus used for generating gas-
phase acetone standards. A known amount of acetone was injected into a calibrated airbag
containing humidified air and allowed to evaporate. The vapor was then released at a fixed
rate and passed through the wetted foam. The acetone concentration of the acetone in the

15 liquid was then measured and correlated to the starting acetone gas phase concentration.
Using this partitioning strategy (that is, wetted foam), it was determined that gas phase
acetone partitioned into the liquid phase to achieve theoretical equilibrium concentration
values for a range of gas flow rates (0.5 to 5.0 L/min.) and gas volumes (> 0.5 L). The
experiment was repeated without foam using a thin liquid film (50 uL) where the acetone

20 gas sample was impinged on the liquid in a fine gas stream. Using this partitioning strategy,
gas phase acetone partitioned into the thin liquid film to achieve near equilibrium
concentration values. These data were then used to devise the appropriate liquid volumes
and conditions (which mimic human breath) for partitioning and quantifying gas-phase
acetone electrochemically using the 3-enzyme system

Figure 23 shows the chronoamperometric response for acetone gas using polyurethane foam
as a means to extract the gas sample. The 3-enzyme system was employed as the base
transducer to convert the partitioned acetone into H2O2 that was then electrochemically
detected using a disk platinum electrode at 350 mV (vs, Ag/AgCl). The current response
30 to gas-phase acetone concentration has excellent sensitivity and linear characteristics over
the entire range tested. Linear regression of the data gave the equation y = 19.181x +
18.787, with an R 2 value of 0.993. The experiment was repeated by partitioning gas-phase
acetone into a liquid film containing the 3-enzyme system. The results are shown in Figure

WO 03/039483 PCT/US02/36028

24 where chronoamperometric current response correlated linearly with the gas-phase
acetone concentration. Linear regression of the data gave the equation y = 26.93 1 x +
28.789, with an R 2 value of 0.9966. These results demonstrate the either strategy for gas-
phase sampling (that is, wetted foam or liquid film) work well as systems to accurately
5 introduce acetone gas samples to the enzyme electrode where they can be quantified
electrochemically.

Biochemical properties of acetone carboxylase. In addition to S-ADH, another acetone-
specific enzyme that may be suitable for use in a biosensor is acetone carboxylase. As

10 mentioned above, acetone carboxylase is a unique enzyme that catalyzes the carboxylation
of acetone to the p-ketoacid acetoacetate in a variety of aerobic and anaerobic bacteria that
are able to grow using acetone as a carbon and energy source. Many of these organisms are
also capable of growth on isopropanol, where as discussed previously, a secondary alcohol
dehydrogenase catalyzes the initial oxidation of isopropanol to acetone, which is then

15 subsequently carboxylated to acetoacetate. Acetone carboxylase can be purified from X
autotrophics strain Py2 via published methods. Acetone carboxylase is a multimeric
enzyme comprised of three polypeptides (19.6 kDa, 78.3 kDa, and 85.3 kDa) arranged in an
CX2P2Y2 quaternary structure and requires MgATP for carboxylation activity. This enzyme
exhibits a V max of 0.206 jimol acetone consumed-min ^mg" 1 protein and apparent K m values

20 of 7.8 jiM (acetone), 122 jiM (ATP), and 4. 1 7 mM (C0 2 plus bicarbonate). Butanone is
also a substrate that is carboxylated at a rate of 40% of that of acetone. Acetone
carboxylase has also been purified from the anaerobic, photoheterotroph Rhodobacter
capsulatus strain B10 and is nearly identical to the Xanthobacter enzyme in terms of
quaternary structure (a 2 p 2 Y2 multimer consisting of 19.5 kDa, 78.6 kDa, and 85.2 kDA

25 polypeptides), kinetic parameters (F max of 0.291 \xmo\ acetone consumed-min'-mg" 1 protein,
AT m of 8.2 \xM for acetone), and amino acid sequence (83% identical).

Schemes proposed for electrochemical detection of the acetone carboxylase reaction.
The ATP-dependent carboxylation of acetone to form acetoacetate, the reaction catalyzed
30 by acetone carboxylase, contains no net redox chemistry for electrochemical detection. To
be useful in a biosensor, it was necessary to develop a coupled enzyme reaction that
provided substrate oxidation or reduction. As shown in Figure 25, acetone carboxylase can
be coupled to P-hydroxybutyrate dehydrogenase where the acetoacetate formed by the

WO 03/039483 PCT/US02/36028

carboxylation reaction is subsequently reduced by this enzyme with concomitant oxidation
of NADH. The acetone-dependent consumption of NADH can then be monitored
electrochemically using the method described above for S-ADH enzyme. Figure 26 shows
a spectrophotometry plot of the rate of NADH oxidation vs. time using this enzyme system.
5 NADH oxidation was observed only after acetone addition and no decrease in the rate was
observed by addition of ethanol (10 mM) indicating that it did not inhibit the reaction. The
enzyme system was highly specific for acetone, but alternative compounds (for example 1-
propanol, 2-propanol, 1-butanol, and 2-butanol) were not active nor did they inhibit acetone
carboxylation activity. Butanone, which normally is a substrate that is carboxylated at a
10 rate of 40% of that of acetone, displayed a rate of about 3 to 4% of acetone using the
coupled enzyme system. This is presumably due to the substrate specificity of the p-
hydroxybutyrate dehydrogenase.

The acetone detection limit of the acetone carboxylase reaction coupled to p-
15 hydroxybutyrate dehydrogenase was investigated using spectrophotometry in a manner
similar to that described above using S-ADH from X. autptrophicus strain Py2. Using
acetone carboxylase and P-hydroxybutyrate dehydrogenase coupled enzyme system, the
NADH consumption response was linear (correlation coefficient = 0.99954) for acetone
concentrations ranging from 0.058 to 2.8 ppm (w/v). The lower end of the acetone
20 detection range (0.058 to 0.29 ppm (w/v)) is within the detection parameters required for
diagnostic breath acetone analysis.

Shown in Figure 27 is an alternative enzyme system that couples acetone carboxylase ATP-
hydrolysis to NADH oxidation. This four component enzyme system, which has been

25 described previously for spectrophotometric detection of acetone carboxylase activity, can
easily be adapted for electrochemical detection (Figure 27). Using this coupled assay,
acetone carboxylase generates AMP as acetone is carboxylated, the AMP that forms is then
converted to two molecules of ADP in a reaction catalyzed by myokinase. The ADP
formed by this reaction is then phosphorylated to ATP by pyruvate kinase which catalyzes

30 the conversion of phosphoenolpyruvate to pyruvate. The pyruvate that forms is then

reduced by lactate dehydrogenase consuming NADH in the process. Although this enzyme
system is more complex, it utilizes the same electrochemical detection method as described
above.

WO 03/039483

PCT/US02/36028

A third means to monitor acetone carboxylase activity electrochemically involves a
modification of the strategy presented in Figure 27, so that ATP hydrolysis is coupled to
H 2 0 2 formation. As shown in Figure 28, in this strategy a pyruvate oxidase is substituted
5 for lactate dehydrogenase; thus, as pyruvate is generated it is oxidized, thereby forming
H2O2. In order to investigate this enzyme system, the enzyme reaction mix (containing
coupling enzymes and reagents) was prepared with acetone carboxylase from Xanthobacter
Py2, along with myokinase, pyruvate kinase, and pyruvate oxidase, in the presence of
horseradish peroxidase (HRP) and electron acceptor dyes to allow monitoring of the

10 reaction system end-product (H 2 0 2 ) spectrophotometrically. Figure 29 illustrates the results
of the coupling reactions, wherein an increase in absorbance was observed, upon addition of
acetone, that corresponded to H 2 0 2 formation. As was the case for the S-ADH-H 2 0 2
generating system, a lag was observed after acetone addition that could be shortened with
further optimization of the reaction conditions. Shown in Figure 30 is a fourth means to

1 5 monitor acetone carboxylase activity electrochemically that involves combining certain
aspects of the strategies diagramed in Figures 27 and 28. Using this strategy, the NAD +
generated from the coupled reaction between acetone carboxylase and p-hydroxybutyrate
dehydrogenase was used by lactate dehydrogenase to oxidize lactate to pyruvate. The
pyruvate thus formed was oxidized by pyruvate oxidase to generate H 2 0 2 which is then

20 detected; electrochemical detection of the H 2 0 2 generated was detected electrochemically
(data not shown).

Also, as an alternative strategy to monitoring acetone-dependent consumption of NAD(P)H,
the acetone mono-oxygenase reaction can be coupled to galactose oxidase to form H202 as
25 shown in Figure 31. Alternatively, acetone mono-oxygenase N AD(P)+ formation can be
coupled to H202 formation by use lactate dehydrogenae and pyruvate oxidase.

Stability of acetone carboxylase. As described above for the S-ADH enzyme, it is
important to devise a formulation for stabilizing acetone carboxylase activity in a dry form
30 for long periods of time at room temperature. Purified acetone carboxylase was lyophilized
and found to retain approximately 90% activity relative to a non-lyophilized control when
rehydrated and assayed within one day of lyophilization (data not shown). However, the
activity decreased over the course of several days, for samples stored at room temperature

WO 03/039483 PCT/US02/36028

and 4°C. As was the case for S-ADH, the presence of trehalose stabilized activity (>90%)
for at least 1 1 1 days in samples held at room temperature. In a separate experiment, the
entire coupling assay reaction mix (that is acetone carboxylase from X. autotrophicus strain
Py2, p-hydroxybutyrate dehydrogenase, NADH, ATP, buffer, potassium acetate, and
5 MgCb) was Iyophilized to determine whether assay components were stable to this
treatment When the assay mix was rehydrated and assayed within one day, 84% of the
activity was retained. When trehalose was included in the reaction mixture that was
Iyophilized, 95% of the activity was retained. A third stability experiment was performed
where acetone carboxylase in the presence of trehalose (20% w/v) was allowed to air dry as
10 opposed to being Iyophilized. In this case, after 24 hours, acetone carboxylase retained a
1 00% of its activity (data not shown). Allowing an enzyme to air dry (instead of
lyophilization) with retention of activity may be an important requirement for screen-
printing technology.

15 Other prospects for incorporating acetone-specific enzymes into an enzyme-based

electrochemical sensor for breath acetone. In some propane-oxidizing bacteria, acetone is
formed as an intermediate that is believed to undergo hydroxylation in an 02-dependent
mono-oxygenase-catalyzed reaction to form acetol (hydroxyacetone). Although this
bacterial enzyme has not been fully characterized, it functions in a manner resembling that

20 of other mono-oxygenase enzymes and systems described to date. Known mono-
oxygenases are typically comprised of multiple enzyme components (2-4 enzyme
components), including a pyridine nucleotide-dependent reductase, an electron transfer
protein (for example ferredoxin), and an active site-containing oxygenase component.
NAD(P)H provides the necessary reductant for O2 activation and incorporation of one

25 oxygen atom into the aliphatic hydrocarbon substrate. Acetone-dependent consumption of
NAD(P)H by an acetone mono-oxygenase reaction could be monitored electrochemically as
described above for secondary alcohol dehydrogenase- and acetone carboxylase-coupled
enzyme systems (Figure 1). In addition, an acetone P450 mono-oxygenase has been
described in mammalian systems. Although this enzyme likely has different biophysical

30 properties (that is different size of subunit molecular masses, amino acid sequences, etc.)
from a bacterially-derived acetone mono-oxygenase, the reaction it catalyzes is suitable for
monitoring acetone in an enzyme-based biosensor, using a strategy parallel to that described
for bacterial acetone mono-oxygenase. Acetol, the product of the acetone mono-oxygenase

WO 03/039483 PCT/USO 2/36028

reaction, has been reported to be a substrate for galactose oxidase (J. Tkac et al., Enzyme
and Microbial Technology (2001) 28: 383-88; M.M. Whittaker and J.W. Whittaker,
Biochemistry (2001) 40:7140-48). Therefore, as an alternative strategy to monitoring
acetone-dependent consumption of NAD(P)H, the acetone mono-oxygenase reaction can be
coupled to galactose oxidase to form H2O2 as shown in Figure 3 1 .

Acetone Signal Amplification Strategies

Amplification of the enzyme electrode signal may be important where acetone
concentrations in tested samples are low. For example, the basal level (in non-fasting
individuals) of acetone in breath is relatively low (0.2 to 0.5 ppm v/v). Amplification can
be accomplished by using enzymes to recycle substrates (futile cycles), thereby magnifying
the output signal. Consequently, a low, sub-saturating level of acetone would yield a
continuous rate of response that would steadily increase the signal intensity over time. The
rate of increase would be dependent on the initial amount of acetone detected, as long as the
concentration is sub-saturating to the enzyme system employed. The biosensor can then be
calibrated by measuring the rates of response and correlating them to the initial acetone
concentration.

In a first embodiment, shown in Figure 32, an amplification scheme uses acetoacetate
decarboxylase. Acetoacetate decarboxylase, found in fermentative microorganisms,
catalyzes the opposite reaction of an acetone carboxylase. Using this scheme, acetone
carboxylase ATP hydrolysis activity is coupled to H 2 0 2 generation as presented in Figure
28, however the addition of acetoacetate decarboxylase recycles acetoacetate back to
acetone, thus creating a system that does not deplete the concentration of acetone. Another
variation of this amplification system is shown in Figure 33, wherein lactate oxidase is
incorporated into the coupled enzyme system presented earlier in Figure 27. As pyruvate is
reduced to lactate, lactate oxidase catalyzes the recycling of lactate back to pyruvate
generating H 2 0 2 in the process that is then detected electrochemically.

A number of different acetone signal amplification strategies can be employed with S-ADH
enzymes. In a first embodiment, illustrated in Figure 34, an S-ADH enzyme is coupled to a
pyrroloquinoline quinone (PQQ)-dependent alcohol dehydrogenase; one examples of such
an enzyme is that isolated from Pseudomonas sp. strain VM15C (Shimao 1986). This

WO 03/039483 PCT/US02/36028

particular PQQ-dehydrogenase has been reported to have specificity to low-molecular-
weight secondary alcohols rather than primary alcohols. In this amplification strategy, an S-
ADH enzyme reduces acetone to isopropanol. The isopropanol thus formed is then
reoxidized to acetone by PQQ-dependent alcohol dehydrogenase. The electrons from this
5 oxidation are transferred directly to the electrode through prosthetic groups of the PQQ
enzyme. Integrated enzyme electrodes involving direct electron transfer have previously
been reported for this class of enzymes, A similar example is shown in Figure 35, in which
an S-ADH enzyme is coupled to a secondary alcohol oxidase (SAO; EC 1.1.3.18). SAOs
have been isolated from a number of different organisms, for example, Pseudomonas sp.

10 See, for example, M. Morita et al., Purification and properties of secondary alcohol oxidase
from a strain of Pseudomonas. Agric. Biol. Chem. 43:1225-35 (1979). Using this strategy,
acetone undergoes recycling, during which it is initially reduced by S-ADH and
subsequently reoxidized by SAO, concurrently producing H2O2 for each turnover of the
substrate cycle. This linear amplification scheme is advantageous in that it is simple (only

15 two enzymes are needed) and takes advantage of the relative ease with which H2O2 can be
monitored.

. This strategy for signal amplification, that is by combining the actions of an alcohol
dehydrogenase and an alcohol oxidase, was investigated by coupling primary alcohol

20 oxidase (AO) to primary alcohol dehydrogenase. Using ethanol as the substrate, and
measuring the increase in H2O2 formation over identical response times, a 7- to 10-fold
increase was observed using the two-enzyme system over AO alone (data not shown). The
signal can be increased further for the two-enzyme system by allowing the response time to
extend for longer periods. This demonstrates that a practical method for amplifying a

25 response to acetone is to couple S-ADH to SAO.

Figure 36 illustrates a third example of an amplification strategy, which enhances the
S-ADH/SAO coupled system by taking advantage of the NAD + produced from the reaction
catalyzed by S-ADH. In this case, NAD + produced from the reduction of acetone to
30 isopropanol is coupled to hydroxybutyrate dehydrogenase (HBDH) where hydroxybutyrate
(loaded in excess) is oxidized to acetoacetate. Acetoacetate is then decarboxylated by
acetoacetate decarboxylase (ACD) to form acetone, thereby increasing the amount of
acetone (by one molecule) that undergoes subsequent recycling. As opposed to the enzyme

WO 03/039483 PCT7US02/36028
system illustrated in Figure 35, which provides linear amplification of an acetone response
(that is for every substrate cycle, the amount of H 2 0 2 produced is doubled), this type of
amplification system would exponentially increase the amount of H2O2 output, allowing
detection of extremely low levels of acetone in a biological sample.

A fourth example is shown in Figure 37, in which an amplification strategy exploits the
preferences of S-ADH enzymes for NADH or NADPH pyridine cofactors. In this case, the
first S-ADH catalyzes the reduction of acetone to isopropanol, while oxidizing NADPH.
The second S-ADH re-oxidizes isopropanol to acetone, while generating NADH that is then
utilized by an NADH-dependent diaphorase (D) to reduce an electron mediator that is then
detected, for example, by oxidation at the electrode. While this system does provide
amplification, a commercial embodiment thereof would preferably employ S-ADH enzymes
having substantially absolute coenzyme specificity, as well as a means to control or actively
monitor the coenzyme concentration(s) and the substrate/product ratios of the chemical
equilibrium.

As described herein, in a preferred embodiment a means is provided for incorporating
acetone-specific enzymes in an electrochemical or non-electrochemical biosensor, thereby
enabling such activities as diagnostic monitoring of acetone on human breath. As discussed
herein, preferable strategies for electrochemical detection of acetone using a combination of
enzymes forming linked enzyme systems include: 1) the secondary alcohol dehydrogenase
(S-ADH)-catalyzed reduction of acetone with concomitant NADPH consumption, and 2) S-
ADH-catalyzed reduction of acetone with concomitant NADH consumption (that is
collectively, S-ADH-catalyzed reduction of acetone with concomitant NAD(P)H
consumption, with NAD(P)H being detected electrochemically); 3) acetone carboxylase
reaction coupled to (for example, P-hydroxybutyrate dehydrogenase) consumption of
NADPH, and 4) acetone carboxylase reaction coupled to (for example, P-hydroxybutyrate
dehydrogenase) consumption of NADH (that is collectively, acetone carboxylase reaction
coupled to consumption of NAD(P)H, with NAD(P)H being detected electrochemically); 5)
acetone carboxylase reaction ATP hydrolysis coupled to NADPH consumption, and 6)
acetone carboxylase reaction ATP hydrolysis coupled to NADH consumption (that is
collectively, acetone carboxylase reaction ATP hydrolysis coupled to NAD(P)H
consumption, with NAD(P)H being detected electrochemically); 7) S-ADH reaction

WO 03/039483 PCT/USO 2/360 28

NADP + formation coupled to H2O2 formation, and 8) S-ADH reaction NAD + formation
coupled to H2O2 formation (that is collectively, S-ADH reaction NAD(P) + formation
coupled to H2O2 formation, with H2O2 being detected electrochemically); 9) acetone
carboxylase reaction ATP hydrolysis coupled to H2O2 formation with H2O2 being detected
5 electrochemically; 1 0) acetone carboxylase reaction coupled to (for example, 0-

hydroxybutyrate dehydrogenase) NADP + formation coupled to H2O2 formation, and 1 1)
acetone carboxylase reaction coupled to (for example, P-hydroxybutyrate dehydrogenase)
NAD + formation coupled to H2O2 formation (that is collectively, acetone carboxylase
reaction coupled to, for example, p-hydroxybutyrate dehydrogenase, NAD(P) + formation

10 coupled to H2O2 formation, with H2O2 being detected electrochemically); and 12) acetone
mono-oxygenase coupled to NADPH oxidation, and 13) acetone mono-oxygenase coupled
to NADH oxidation (that is collectively, acetone mono-oxygenase coupled to NAD(P)H
oxidation). The invention is not limited to these particular enzyme systems, and any
acetone-specific enzyme may be employed in the enzyme systems of the invention.

15 Moreover, any electrochemically detectable cofactors and by-products suitable for coupling
with acetone-specific enzymes may be employed.

Example 5

An acetone biosensor according to the present invention may be utilized to track the release
20 of acetone formed by metabolism of a "tag" compound administered as part of a drug

formulation or other administered or implanted composition. In a preferred embodiment,
acetoacetic acid and/or any of its pharmaceutical^ acceptable salts and pharmaceutical^
acceptable esters or amides may be used as a tag to track the release of compositions
administered to, or implanted into, a living organism or a group of living organisms, for
25 example, an environmental sample or a cell culture or colony. Pharmaceutical^ acceptable
base addition salts of acetoacetic acid include alkali metals, such as sodium, potassium and
lithium salts; alkaline earth metals, such as calcium and magnesium; transition metals, such
as zinc, iron and copper; carbonate, bicarbonate, ammonium, alkylammonium, alkylamine,
alkanolamine and hydroxy alkamine salts. Also suitable are alcohol esters. Preferably,
30 sodium or potassium salts of acetoacetic acid are used. A preferred ester is ethyl ester.
Acetoacetic acid, as well sodium and potassium salts thereof, are solids at room
temperature, whereas ethyl acetoacetate is a liquid at room temperature. Salts of acetoacetic

WO 03/039483 PCT/US02/36028

acid may be administered for tagging purposes at a dosage of about 1 0 mg to about 200 mg,
preferably in a range from about 10 mg to about 170 mg.

During p-oxidation of fatty acids, two molecules of acetyl CoA are hydrolyzed from a fatty
5 acid chain with each oxidation cycle, during which ATP is generated for energy needs. The
acetyl CoA so produced may be metabolized to carbon dioxide, or it may be utilized in the
formation of ketone bodies via conversion into acetoacetate in the liver of a mammal. The
acetoacetate produced in this manner may serve as an alternative to glucose as an energy
source, particularly during periods of starvation. Acetoacetate produced by the liver is

10 released into the bloodstream for use as fuel, particularly by the brain and heart muscle.
Free acetoacetic acid infused into the bloodstream of an animal has been shown to induce
increases in alanine and glutamine levels, possibly due to a stimulation of muscular output
of these amino acids, while decreases in blood glucose levels have been noted. In general,
acetone is rapidly cleared from mammalian systems via metabolism/exhalation and

15 excretion in urine. Acetoacetate spontaneously decarboxylates to form acetone and CO2,
and the odor of acetone is noticeable in the breath of individuals having high blood levels of
acetoacetate. Exhalation as C0 2 and unchanged acetone is the primary route of acetone
elimination from the body. The amount of unchanged acetone exhaled is related to
concentrations of acetone in the body. Both urinary and exhaled acetone can be detected in

20 biological samples using an acetone-specific biosensor.

Thus, an acetone-specific biosensor may be employed to detect the release of acetone
derived from the acetoacetate tag and thereby track the release of an administered
therapeutic compound or implanted composition, or the biodegradation and/or bio-erosion
25 of the implanted composition.

In a preferred embodiment, the administered formulation will contain the acetoacetic acid,
salt, ester, or amide in admixture with a primary substance whose bio-release is desired.
Examples of such primary substances include, but are not limited to: pharmaceutically
30 acceptable active ingredients, biopharmaceuticals (for example, proteins such as antibodies,
hormones, enzymes, serums, and vaccines; gene therapies), and other bioactive substances
such as dietary supplements (for example, vitamins, minerals, herbal supplements, and
nutraceuticals).

WO 03/039483 PCT/USO 2/36028

Pharmaceutical active ingredients include, but are not limited to: analgesics, anesthetics,
antacids, anthelmintics, antibiotics, anticoagulants, anticonvulsants, antidepressants, anti-
emetics, antifungals, antihistamines, antihypertensives, anti-infectives, antiinflammatories,
5 antimanic agents, antimicrobials, antineoplastic agents, antiparasitics, antiprotozoals,

antipsychotics, antipyretics, antiseptics, antitussives, antivirals, autonomic agents (such as
anticholinergics, sympathomimetics, sympatholytics, parasympathomimetics,
parasympatholytics), bronchodilators, cardiovascular drugs, cathartics, chemotherapeutic
agents, coagulants, contraceptives, depressants, diuretics, expectorants, hematopoietic
10 agents, hypnotics, immunomodulators, psychopharmacologic agents, sedatives, stimulants,
tranquilizers, vasoconstrictors, and vasodilators.

In a preferred embodiment, the implanted composition will be a solid material into which
the acetoacetic acid, salt, amide, or ester, or formulation therewith has been loaded or

1 5 imbedded, for example, as powdered- or granulated-particles or liquid- or solution-droplets,
distributed throughout the bulk of the solid material and/or among the interstices thereof.
The particles or droplets may consist solely of the acetoacetic acid, salt, ester, or amide or
the solution thereof, or they may incorporate additional substance(s), including, but not
limited to, the above-mentioned primary substances, and pharmaceutically acceptable

20 excipients, adjuvants, diluents, and/or carriers. The solid material may be rigid, or it may be
a plastic, rubber, gel and/or foam material, and it may take any form, for example, lump,
block, bar, pellet, sheet, film, membrane, fiber, mat, mesh, or matrix form.

Tagging a pharmaceutical with acetoacetate, or a derivative thereof, permits easy detection
25 of subject compliance with prescribed therapeutic regimes by virtue of detecting breath

acetone using an acetone specific enzyme system according to the present invention. Thus,
subjects mentally impaired due to age or disease, mentally ill subjects, or any other poorly
compliant subject can be monitored for drug dosages in a noninvasive manner. By
integrating an acetone-specific biosensor with an analysis device linked to, for example, the
30 Internet, instant results for home monitoring of a subject can be obtained. In this manner,
acetone-specific biosensors may be employed to ensure health care services are being
delivered to a given subject, to test the bio-release of drug formulations in clinical test
settings, and to test the release of delayed or controlled release drug formulations over time.

WO 03/039483 PCTAJS02/36028

Acetone-specific biosensors may also be useful in a biotic environment, such as part of a
medical device, implant, and the like. Such devices and implants include drug delivery
implants, cosmetic implants, prosthetic implants, and bionic implants.

5 As noted above, acetoacetic acid is a solid at room temperature. Thus, acetoacetic acid and
its pharmaceutically acceptable salts and pharmaceutical ly acceptable esters and amides that
are solid at room temperature are appropriate for use in, for example, powdered, granular,
pill, tablet, capsule, implant, and solution formulations. The pharmaceutically acceptable
salts and pharmaceutically acceptable esters and amides of acetoacetic acid that are liquid at

10 room temperature are appropriate for use in, for example, liquid formulations (for example,
elixirs, syrups, drops, and liquid capsules) and semirliquid formulations (for example,
pastes, creams, and ointments). Preferred solid formulations include pill, tablet, and capsule
formulations, more preferably non-chewable pills, tablets, and capsules, and also include
implant formulations. Preferred solution, liquid, and semi-liquid formulations include

15 injectable and infusible formulations, more preferably parenteral injectable and infusible
formulations, and also include implant formulations.

The term "alkyl" when used in regard to pharmaceutically acceptable salts, esters, and
amides is defined herein as "aliphatic. "Preferred "alkyl" groups include CI -CIO aliphatic
20 groups, more preferably unsaturated C 1 -C 1 0 aliphatic groups, still more preferably

unsaturated C1-C6 aliphatic groups. Even more preferably, where a straight-chain alkyl
group is used, it is preferably an unsaturated C1-C4 aliphatic group.

An example of drug tagging via use of an acetone-specific biosensor would be monitoring
25 the intake of lithium carbonate by a subject diagnosed with bipolar disorder. Fairly strict
titrations of lithium are needed to achieve reasonable plasma levels of for effectiveness
without incurring excess levels, which can be fatal. Thus, a capsule of 300 mg lithium
carbonate may be reformulated to contain 200 mg of sodium acetoacetate, which is
administered to a subject in a standard fashion (for example, four times per day). Subject
30 compliance can then be monitored using an acetone-specific biosensor to detect acetone
released from the acetoacetate tag. By interfacing the biosensor with the Internet, the
subject's physician can log the daily acetone levels measured over a period of time. Serum
lithium levels can be simultaneously measured to assign a correlation between acetone

WO 03/039483 PCT/US02/36028

measurements and bioavailability of lithium. Thus, a biosensor according to the present
invention can be applied to reduce or eliminate the need for invasive (e.g., serum) testing
for lithium levels.

5 Alternatively, the acetone tag may be co-administered with the pharmaceutical^ active
compound in the form of a placebo, instead of being formulated with the compound.
Methods of infusing acetoacetate into human subjects are known in the art. Subjects can be
infused over a several hour period, for example, for three hours, with Na acetoacetate or
free acetoacetic acid administered at approximately 20 |imol/kg/min. Constant perfusion of

10 Na acetoacetate at a rate of 1 .36 mmol/min. for three hours has been shown to cause an
increase in blood ketone body concentration to 3 |imol/mL at the end of perfusion.
Constant infusions of 3- 14 C-Iabeled acetoacetate, sodium salt (for example, infused at a rate
of 0.68-0.88 nanocuries per kilogram per minute (mjiCi/kg/min.)) permit the tracking of
respiratory efflux of l4 C02. Thus, perfusion of a tagged acetoacetate together with a drug of

15 interest would provide a means for correlating acetone production with blood bioavailability
of a drug of interest.

Also contemplated is a kit for detecting acetone in a sample comprising the acetone-specific
enzyme system separate from, or contained within, a biosensor, together with instructions
20 for use. For example, the acetone-specific enzyme system may be adhered to or otherwise
disposed on a disposable test strip that may be inserted into a housing, the housing having a
port into which a biological sample may be introduced. Alternatively, the biosensor
housing and the acetone-specific enzyme system may be fashioned to be a single,
disposable unit.

Given the limitations of the known means of detecting acetone in a biological sample, the
inventive acetone-specific enzyme systems and biosensors containing these enzyme systems
offer the advantages of being more sensitive to low concentrations of acetone, more specific
for acetone, and less likely to give false readings due to interference from impurities.
30 Therefore, the inventive acetone-specific biosensors are suitable for use outside the
laboratory, and may be portable. Portable acetone-specific biosensors having high
sensitivity for acetone would be convenient for monitoring acetone levels in subjects in a

WO 03/039483 PCT/US02/36028

variety of settings, yet are expected to exhibit at least the level of accuracy attributed to
traditional nitroprusside tests for acetone in body fluids.

The various signal amplification schemes disclosed herein may also be employed in systems
for detection components other than acetone, by substituting for the acetone-specific
enzyme, an different enzyme capable of generating the same cofactor (e.g., NAD(P)H or
NAD(P)+) or reaction by-product as the acetone-specific enzyme.

The concepts and data presented here provide avenues for exploiting an enzyme-based
biosensor specific for acetone detection. While the invention has been illustrated by the
preferred embodiments described herein, those skilled in the art will appreciate that various
other modifications will be apparent and can be readily made by such artisans without
departing from the spirit and scope of the invention. Accordingly, it is not intended that the
scope of the appended claims be limited to the preferred embodiments described herein, but
broadly construed the full extent supported by the disclosure as a whole. The disclosures of
all references cited are hereby incorporated herein by reference.

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WHAT IS CLAIMED IS:

PCT/US02/36028

1 . An acetone-specific enzyme system including an enzyme that selectively targets
acetone as a substrate, coupled to a detectable signal mediator.

2. The acetone-specific enzyme system according to claim 1 , wherein the acetone-
specific enzyme is any one of an acetone mono-oxygenase, an acetone carboxylase, or a
secondary alcohol dehydrogenase.

10 3. The acetone-specific enzyme system according to claim 1 , wherein the acetone-

specific enzyme system is coupled to a signal mediator detectable by electrochemical or
photometric means.

4. The acetone-specific enzyme system according to claim 3, wherein the acetone-
15 specific enzyme system is any one of acetone carboxylase product formation coupled to
NAD(P)H oxidation, acetone carboxylase ATP-hydrolysis coupled to NAD(P)H oxidation,
acetone carboxylase ATP hydrolysis coupled to H 2 0 2 formation, secondary alcohol
dehydrogenase (S-ADH) with concomitant NAD(P)H oxidation, acetone carboxylase
coupled to (J-hydroxybutyrate dehydrogenase catalyzed NAD(P) + formation coupled to
20 H 2 0 2 production, S-ADH catalyzed NAD(P) + formation coupled to H 2 0 2 production,
acetone mono-oxygenase coupled to NAD(P)H oxidation, or acetone mono-oxygenase
coupled to H 2 0 2 production, or acetone monooxygenase-catalyzed NAD(P)+ formation
coupled to H202 formation.

25 5. The acetone-specific enzyme system according to claim 4, wherein the secondary

alcohol dehydrogenase is obtained from a mammal or from a microorganism that is any one
of the aerobic bacteria, anaerobic bacteria, yeasts, fungi, or methanogenic Archaea.

6. The acetone-specific enzyme system according to claim 5, wherein the secondary
30 alcohol dehydrogenase is isolated from a species of any one of Thermocmaerobium,
Xanthobacter, Pseudomonas, Rhodococcus, or Mycobacterium.

118

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7. The acetone-specific enzyme system according to claim 5, wherein the secondary
alcohol dehydrogenase is isolated from Xanthobacter autotrophics strain Py2.

8. The acetone-specific enzyme system according to claim 4, wherein the acetone
carboxylase is isolated from a species of any one of Xanthobacter, Rhodobacter, or
Rhodococcus.

9. The acetone-specific enzyme system according claim 4, wherein the signal
mediator is a member of any one of organic cofactors, inorganic cofactors, indicators,
electron transfer mediators, photometric mediators, enzyme reaction by-products, or
combinations thereof.

10. The acetone-specific enzyme system according to claim 9, wherein a signal from
the electrochemically detectable signal mediator is linearly or exponentially amplified by
recycling enzyme substrates to magnify electrochemical signal output.

1 1. A method for using an acetone-specific enzyme system according to claim 1,
wherein said method involves

a) combining

i) at least one pharmaceutical compound, with

ii) at least one tag selected from the group consisting of acetoacetate and
pharmaceutically acceptable acetoacetate salts, esters, and amides, to form a tagged
pharmaceutical compound;

b) administering the tagged compound to a subject; and

c) thereafter, detecting release of the tag into the subject's bloodstream via
measurement, using the acetone-specific enzyme system, of the acetone concentration in a
biological sample from the subject, wherein the acetone is present in the subject's biological
sample due to degradation of the tag, forming the acetone;

d) correlating the acetone concentration measured from the biological sample with
release of the pharmaceutical compound from the administered tagged compound; and

e) optionally, repeating steps c) and d) to determine a rate of release of the
pharmaceutical compound from the administered tagged compound.

119

WO 03/039483 PCT/US02/36028

12. A method for using an acetone-specific enzyme system according to claim 1,
wherein said method involves

a) combining

i) at least one tag selected from the group consisting of acetoacetate and
pharmaceutical^ acceptable acetoacetate salts, esters, and amides, with

ii) a biocompatible composition, to form a biocompatible implant or device;

b) placing the biocompatible implant or device in a subject;

c) thereafter, detecting release of acetoacetate into the subject's bloodstream via
measurement, using the acetone-specific enzyme system, of the acetone concentration in a
biological sample from the subject, wherein the acetone is present in the subject's biological
sample due to degradation of the tag, forming the acetone;

d) correlating the acetone concentration measured from the biological sample with
degradation or release of the biocompatible composition from the biocompatible implant or
device; and

e) optionally, repeating steps c) and d) to determine a rate of degradation or release
of the biocompatible implant or device.

13. A method for using an acetone-specific enzyme system according to claim 1,
wherein said method involves

tagging a biocompatible implant or device by combining at least one member
selected from the group consisting of acetoacetate, an acetoacetate salt, an acetoacetate ester
and an acetoacetate amide with a biocompatible composition to form a biocompatible
implant or device;

placing the biocompatible implant or device in a subject; and

measuring release of acetoacetate from the biocompatible implant or device.

14. A method for detecting acetone in a biological sample, said method involving:
introducing a biological containing acetone to a biosensor comprising at least one

acetone-specific enzyme system that utilizes acetone as a substrate; and

detecting products produced upon interaction between the acetone and acetone-
specific enzyme system.

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15. The method according to claim 14, wherein detection is achieved by
electrochemical, photometric, or calorimetric means.

1 6. The method according to claim 14, wherein said method farther includes
facilitating electrochemical transduction of the at least one acetone-specific enzyme

system and the acetone in the biological sample via an electrochemically treated electrode;
and

electrochemically detecting a product resulting from a reaction between the at least
one acetone-specific enzyme system and the acetone in the biological sample.

17. The method according to claim 14, wherein the at least one acetone-specific
enzyme system includes an enzyme that is any one of acetone mono-oxygenase, acetone
carboxylase, or secondary alcohol dehydrogenase.

1 8. The method according to claim 1 7, wherein the at least one acetone-specific
enzyme system includes a member that is any one of acetone carboxylase product formation
coupled to NAD(P)H oxidation, acetone carboxylase ATP-hydrolysis coupled to NAD(P)H
oxidation, acetone carboxylase ATP hydrolysis coupled to H2O2 formation, secondary
alcohol dehydrogenase (S-ADH) with concomitant NAD(P)H oxidation, acetone
carboxylase coupled to p-hydroxybutyrate dehydrogenase catalyzed NAD(P) + formation
coupled to H 2 0 2 production, S-ADH catalyzed NAD(P) + formation coupled to H 2 0 2
production, or acetone mono-oxygenase coupled to NAD(P)H oxidation.

19. The method according to claim 18, wherein the biological sample is in vapor

form.

20. The method according to claim 1 9, wherein said method involves
electrochemically detecting acetone in the vapor sample at a level of 0.2ppm to lOppm.

21 . A biosensor for detecting acetone in a biological sample containing:

at least one acetone-specific enzyme system comprising an enzyme that selectively
targets acetone as a substrate, coupled to a detectable signal mediator; and

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WO 03/039483 PCT/USO 2/36028

a means for detecting a product resulting from a reaction between the at least one
acetone-specific enzyme system and acetone in the biological sample.

22. The biosensor according to claim 21, further including a housing having a port
5 for introducing the biological sample to the acetone-specific enzyme system.

23. The biosensor according to claim 21, wherein the acetone-specific enzyme
system is present in a dried or lyophilized form until contacting the biological sample.

10 24. The biosensor according to claim 21 , wherein the detection is achieved via

electrochemical, photometric, or calorimetric means.

25. An electrochemical biosensor according to claim 24, wherein the at least one
acetone-specific enzyme system includes an enzyme that is any one of acetone rnono-

15 oxygenase, acetone carboxylase, or secondary alcohol dehydrogenase.

26. The electrochemical biosensor according to claim 25, wherein the at least one
acetone-specific enzyme system is a member that is any one of acetone carboxylase product
formation coupled to NAD(P)H oxidation, acetone carboxylase ATP-hydrolysis coupled to

20 NAD(P)H oxidation, acetone carboxylase ATP hydrolysis coupled to H2O2 formation,

secondary alcohol dehydrogenase (S-ADH) with concomitant NAD(P)H oxidation, acetone
carboxylase coupled to P-hydroxybutyrate dehydrogenase catalyzed NAD(P) + formation
coupled to H 2 0 2 production, S-ADH catalyzed NAD(P) + formation coupled to H 2 0 2
production, acetone mono-oxygenase coupled to NAD(P)H oxidation, acetone mono-

25 oxygenase coupled to H2O2 production, or acetone monooxygenase-catalyzed NAD(P)+
formation coupled to H202 formation.

27. An electrochemical biosensor according to claim 25, wherein the acetone-
specific enzyme system is storage stabilized by the presence of trehalose.

28. An electrochemical biosensor according to claim 25, wherein the biological
sample is either in liquid or vapor form.

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29. An electrochemical biosensor according to claim 25, further including either
means for generating, from the electrochemical response that occurs during acetone
detection, a qualitative or quantitative result, or means for interfacing said biosensor with
said means for generating a qualitative or quantitative result.

30. An electrochemical biosensor according to claim 25, having at least two analyte
detection electrodes that are grouped in a sequentially arranged cluster along a sample
detection pathway, wherein at least one of said analyte detection electrodes is an acetone-
specific electrode operably comprising an acetone-specific enzyme system.

31 . A method of using the biosensor according to claim 20 to monitor a subject's
medical condition, wherein the method involves

a) obtaining at least one biological sample from the subject and introducing the
biological sample to the biosensor;

b) measuring acetone concentration in the biological sample(s) via detecting acetone
interactions with the acetone-specific enzyme system of the biosensor;

c) optionally, interfacing the biosensor with a computer network to transmit the
measurement(s); and

d) correlating the acetone concentration measurement(s) with the subject's medical
condition.

32. The method according to claim 31, wherein the medical condition being
monitored is diabetes or weight loss.

33. A kit for detecting acetone in a sample, the kit comprising at least one acetone-
specific enzyme system which includes an enzyme that selectively targets acetone as a
substrate, coupled to a detectable signal mediator; and a housing for the at least one
acetone-specific enzyme system.

34. The kit according to claim 33 further containing a disposable test strip upon
which the acetone-specific enzyme system is disposed.

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WO 03/039483 PCT/US02/36028

35. The kit according to claim 34, wherein the strip is separate from the housing
such that the strip may be inserted into the housing, the housing having a port into which the
sample may be introduced.

5 36. The kit according to claim 34, wherein the strip and the housing for the strip are

fashioned as a single, disposable unit, the housing having a port into which the sample may
be introduced.

37. A disposable test strip upon which an acetone-specific enzyme or acetone-
10 specific enzyme system is disposed.

38. The strip according to Claim 37, wherein the acetone-specific enzyme is any one
of an acetone mono-oxygenase, an acetone carboxylase, or a secondary alcohol
dehydrogenase.

39. A protein obtained from Xanthobacter autotrophic™ Py2 (ATCC PTA-4779),
having NAD + -dependent secondary alcohol dehydrogenase activity, having the ability to
reduce acetone to isopropanol, and having specific activity for ketones and secondary
alcohols;

20 (A) said protein (1) having, for the oxidation of isopropanol to acetone, a pH optimum of
approximately 7.8, and (2) having, for the oxidation of alcohols, an average specific activity
ratio for secondary-to-primary alcohols of at least 50:1 when tested at pH 7.8 under
equivalent conditions individually with C3-C5 straight chain secondary alcohols and with
C2-C5 straight chain primary alcohols;

25 (B) said protein having, for the reduction of acetone to isopropanol, (1) a pH optimum of
approximately 6.2, (2) an apparent K m of approximately 144 ± 18 (iM, (3) an apparent K max
of approximately 43.4 ± 1.2 \ixnol acetone reduced min^ mg" 1 protein, (4) an apparent Acai of
approximately 30.4 sec" 1 , (5) an apparent hJK m of approximately 2.1 x 10 5 , and (6) a K m
for NADH of approximately 5.1 ± 0.4 \xM; and

30 (C) said protein comprising at least one polypeptide molecule, (1) said polypeptide

molecule having (a) a molecular mass of approximately 37.1 kDa as determined by mass
spectrometry, (b) a pi of approximately 7.4 as determined by isofocusing electrophoresis,
and (c) a tetradecameric N-terminal amino acid sequence of SEQ ID NO:7, and (2) said

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WO 03/039483 PCT/US02/36028

polypeptide molecule being capable of being degraded to form fragments having the amino
acid sequences of SEQ ID NO:8 to SEQ ID NO: 19.

40. A polypeptide molecule obtained from the protein according to Claim 37, (1)
said polypeptide molecule having (a) a molecular mass of approximately 37.1 kDa as
determined by mass spectrometry, (b) a pi of approximately 7.4 as determined by
isofocusing electrophoresis, and (c) a tetradecameric N-terminal amino acid sequence of
SEQ ID NO:7, and (2) said polypeptide molecule being capable of being degraded to form
fragments having the amino acid sequences of SEQ ID NO:8 to SEQ ID NO: 19.

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SEQUENCE LISTING

<110> Dow Global Technologies Inc.
Cranley, Paul E.
5 Allen, Jeffrey R.

Subramanian, Venkiteswaran
Miller, Theodore E.
Strickland, Alan D.
Danowski, Kristine L.
10 Mclntyre, James A.

<120> Enzyme-Based System and Sensor for Measuring Acetone
<130> 61776A

<150> US 60/332,349
<151> 2001-11-09

<160> 19

<170> Microsoft Word 97-SR2

<210> 1
<211> 776
25 <212> PRT

<213> Xanthobacter autotrophics Py2

<220>

<221> DOMAIN
30 <222> 1. .776

<223> Acetone carboxylase, alpha subunit

<400> 1

Met Asn Val Thr Val Asp Gin Ser Thr Leu Ala Gly Ala Thr Arg Gly
.35 1 5 10 15

lie Val Arg Gly Gly Glu Thr Leu Lys Glu His Arg Asp Arg Leu Met
20 25 30

40 Ala Ala Thr Lys Ala Thr Gly Arg. Tyr Ala Gly Leu Lys Thr Leu Glu
35 40 45

Leu Arg Glu Arg Glu Pro He Leu Tyr Asn Lys Leu Phe Ser Arg Leu
50 55 60

Arg Ala Gly Val Val Asp Ala Arg Glu Thr Ala Lys Lys He Ala Ala
65 70 75 80

Ser Pro He Val Glu Gin Glu Gly Glu Leu Cys Phe Thr Leu Tyr Asn
50 85 90 95

Ala Ala Gly Asp Ser Leu Leu Thr Ser Thr Gly He He He His Val
100 105 HO

55 Gly Thr Met Gly Ala Ala He Lys Tyr Met He Glu Asn Asn Trp Glu
115 120 -125

Ala Asn Pro Gly Val His Asp Lys Asp He Phe Cys Asn Asn Asp Ser
130 135 140

102

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Leu He Gly Asn Val His Pro Cys Asp He His Thr He Val. Pro He
145 150 155 160

Phe Trp Glu Gly Glu LeU He Gly Trp Val Gly Gly Val Thr His Val
5 165 170 175

He Asp Thr Gly Ala Val Gly Pro Gly Ser Met Ala Thr Gly Gin Val
180 185 190

10 Gin Arg Phe Gly Asp Gly Tyr Ser He Thr Cys Arg Lys Val Gly Ala
195 200 205

Asn Asp Thr Leu Phe Arg Asp Trp Leu His Glu Ser Gin Arg Met Val
210 215 220

Arg Thr Thr Arg Tyr Trp Met Leu Asp Glu Arg Thr Arg He Ala Gly

225 230 235 240

Cys His Met He Arg Lys Leu Val Glu Glu Val Val Ala Glu Glu Gly
20 245 250 255

lie Glu Ala Tyr Trp Lys Phe Ala Tyr Glu Ala Val Glu His Gly Arg
260 265 270

25 Leu Gly Leu Gin Ala Arg He Lys Ala Met Thr He Pro Gly Thr Tyr
275 280 285

Arg Gin Val Gly Phe Val Asp Val Pro Tyr Ala His Glu Asp Val Arg

290 295 300

Val Pro Ser Asp Phe Ala Lys Leu Asp Thr He Met His Ala Pro Cys

305 310 315 320

Glu Met Thr He Arg Arg Asp Gly Thr Trp Arg Leu Asp Phe Glu Gly
325 330 335

Ser Ser Arg Trp Gly Trp His Thr Tyr Asn Ala His Gin Val Ser Phe
340 345 350

40 Thr Ser Gly He Trp Val Met Met Thr Gin Thr Leu He Pro Ser Glu
355 360 365

Met He Asn Asp Gly Ala Ala Tyr Gly Thr Glu Phe Arg Leu Pro Lys

370 375 380

Gly Thr Trp Met Asn Pro Asp Asp Arg Arg Val Ala Phe Ser Tyr Ser

385 390 395 400

Trp His Phe Leu Val Ser Ala Trp Thr Ala Leu Trp Arg Gly Leu Ser

50 405 410 415

Arg Ser Tyr Phe Gly Arg Gly Tyr Leu Glu Glu Val Asn Ala Gly Asn

420 425 430

55 Ala Asn Thr Ser Asn Trp Leu Gin Gly Gly Gly Phe Asn Gin Tyr Asp

435 440 " " 445

Glu He His Ala Val Asn Ser Phe Glu Cys Ala Ala Asn Gly Thr Gly
450 455 460

103

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Ala Thr Ala Val Gin Asp Gly Leu Ser His Ala Ala Ala He Trp Asn
465 470 475 480

Pro Glu Gly Asp Met Gly Asp Met Glu He Trp Glu Leu Ala Glu Pro
485 490 * 495

Leu Val Tyr Leu Gly Arg Gin He £ys Ala Ser Ser Gly Gly Ser Gly
500 505 510

Lys Tyr Arg Gly Gly Cys Gly Phe Glu Ser Leu Arg Met Val Trp Asn
515 520 525

Ala Lys Asp Trp Thr Met Phe Phe Met Gly Asn Gly His He Ser Ser
530 535 540

Asp Trp Gly Leu Met Gly Gly Tyr Pro Ala Ala Ser Gly Tyr Arg Phe
545 550 555 * 560

Ala Ala His Lys Thr Asn Leu Lys Glu Leu He Ala Ser Gly Ala Glu
565 570 575

He Pro Leu Gly Gly Asp Thr Asp Pro Glu Asn Pro Thr Trp Asp Ala
580 585 590

Met Leu Pro Asp Ala Gin He Lys Arg Asp Lys Gin Ala He Thr Thr
595 600 605

Glu Glu Met Phe Ser Asp Tyr Asp Leu Tyr Leu Asn Tyr Met Arg Gly
610 615 620

Gly Pro Gly Phe Gly Asp Pro Leu Asp Arg Glu Pro Gin Ala Val Ala
625 630 635 640

Asp Asp He Asn Gly Gly Tyr Val Leu Glu Arg Phe Ala Gly Glu Val
645 650 655

Tyr Gly Val Val Val Arg Lys Gly Ala Asp Gly Gin Tyr Gly Val Asp
660 665 670

Glu Ala Gly Thr Ala Ala Ala Arg Ala Gin lie Arg Lys Asp Arg Leu
675 680 685

Ala Lys Ser Val Pro Val Ser Glu Trp Met Lys Gly Glu Arg Glu Lys
690 695 700

He Leu Ala Lys Asp Ala Gly Thr Gin Val Arg Gin Met Phe Ala Ala
705 710 715 720

Ser Phe Lys Leu Gly Pro Arg Phe Glu Lys Asp Phe Arg Thr Phe Trp

Ser Leu Pro Asp Ser Trp Thr Leu Pro Glu Glu Glu He Gly Val Pro
740 745 750

Thr Tyr Gly Ser Arg Tyr Ser Met Asp He Ser Glu Leu Pro Asp Val
755 760 7 65

His Thr Val Gin Phe Val Glu Glu

725

730

735

770

775

104

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<210> 2

<211> 717

<212> PRT

<213> Xanthobacter autotrophics Py2

<220>

<221> DOMAIN

<222> 1. .717

<223> Acetone carboxylase, beta subunit

<400> 2

Met Asn Val Pro Val Gly His Leu Arg Asn Val Gin Val Leu Gly lie
1 5 10 15

15 Asp Ala Gly Gly Thr Met Thr Asp Thr Phe Phe Val Asp Gin Asp Gly
20 25 30

Asp Phe Val Val Gly Lys Ala Gin Ser Thr Pro Gin Asn Glu Ala Leu
35 40 45

Gly Leu lie Ala Ser Ser Glu Asp Gly Leu Ala Asn Trp Gly Met Ser
50 55 60

Leu His Glu Ala Leu Ala Gin Leu Gin Thr Gly Val Tyr Ser Gly Thr
25 65 70 75 80

Ala Met Leu Asn Arg Val Val Gin Arg Lys Gly Leu Lys Cys Gly Leu
85 90 95

30 He Val Asn Arg Gly Met Glu Asp Phe His Arg Met Gly Arg Ala Val
100 105 110

Gin Ser His Leu Gly Tyr Ala Tyr Glu Asp Arg He His Leu Asn Thr
115 120 125

His Arg Tyr Asp Pro Pro Leu Val Pro Arg His Leu Thr Arg Gly Val
130 135 140

Val Glu Arg Thr Asp Met He Gly Thr Gin Val He Pro Leu Arg Glu
40 145 150 155 160

Asp Thr Ala Arg Asp Ala Ala Arg Asp Leu He Ala Ala Asp Ala Glu
165 170 175

45 Gly He Val He Ser Leu Leu His Ser Tyr Lys Asn Pro Glu Asn Glu
180 185 " 190

Arg Arg Val Arg Asp He Val Leu Glu Glu Val Glu Lys Ser Gly Lys
195 200 205

Lys He Pro Val Phe Ala Ser Ala Asp Tyr Tyr Pro Val Arg Lys Glu
210 215 ~ 220

Thr His Arg Thr Asn Thr Thr He Leu Glu Gly Tyr Ala Ala Glu Pro
55 225 230 235 240

Ser Arg Gin Thr Leu Ser Lys He Ser Asn Ala Phe Lys Glu Arg Gly
245 250 255

60 Thr Lys Phe Asp Phe Arg Val Met Ala Thr His Gly Gly Thr He Ser
260 265 " 270

105

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Trp Lys Ala Lys Glu Leu Ala Arg Thr He Val Ser Gly Pro He Gly

275 280 285

5 Gly Val He Gly Ala Lys Tyr Leu Gly Glu Val Leu Gly Tyr Lys Asn

290 295 300

He Ala Cys Ser Asp He Gly Gly Thr Ser Phe Asp Val Ala Leu He

305 310 315 320

Thr Gin Gly Glu Met Thr He Lys Asn Asp Pro Asp Met Ala Arg Leu
325 330 335

Val Leu Ser Leu Pro Leu Val Ala Met Asp Ser Val Gly Ala Gly Ala
15 340 345 350

Gly Ser Phe He Arg Leu Asp Pro Tyr Thr Arg Ala He Lys Leu Gly
355 360 " 365

20 Pro Asp Ser Ala Gly Tyr Arg Val Gly Val Cys Trp Lys Glu Ser Gly
370 . 375 380

He Glu Thr Val Thr He Ser Asp Cys His Met Val Leu Gly Tyr Leu
385 390 395 400

Asn Pro Asp Asn Phe Leu Gly Gly Ala Val Lys Leu Asp Arg Gin Arg
405 410 ^ 415

Ser Val Asp Ala He Lys Ala Gin He Ala Asp Pro Leu Gly Leu Ser
30 420 425 430

Val Glu Asp Ala Ala Ala Gly Val He Glu Leu Leu Asp Ser Asp Leu
435 440 445

35 Arg Asp Tyr Leu Arg Ser Met He Ser Gly Lys Gly Tyr Ser Pro Ala
450 455 460

Ser Phe Val Cys Phe Ser Tyr Gly Gly Ala Gly Pro Val His Thr Tyr

465 470 475 480

Gly Tyr Thr Glu Gly Leu Gly Phe Glu Asp Val He Val Pro Ala Trp

4 85- 4 90 4 95

Ala Ala Gly Phe Ser Ala Phe Gly Cys Ala Ala Ala Asp Phe Glu Tyr

45 500 505 510

Arg Tyr Asp Lys Ser Leu Asp He Asn Met Pro Thr Glu Thr Pro Asp

.515 520 525

50 Thr Asp Lys Glu Lys Ala Ala Ala Thr Leu Gin Ala Ala Trp Glu Glu
530 535 540

Leu Thr Lys Asn Val Leu Glu Glu Phe Lys Leu Asn Gly Tyr Ser Ala
545 550 555 ~ 560

Asp Gin Val Thr Leu Gin Pro Gly Tyr Arg Met Gin Tyr Arg Gly Gin
565 570 575

Leu Asn Asp Leu Glu He Glu Ser Pro Leu Ala Gin Ala His Thr Ala
60 580 585 590

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Ala Asp Trp Asp Gin Leu Thr Asp Ala Phe Asn Ala Thr Tyr Gly Arg
595 600 605

" ValTyr Ala Ala Ser Ala Arg Ser Pro Glu Leu Gly Tyr Ser Val Thr
5 610 615 620

Gly Ala lie Met Arg Gly Met Val Pro lie Pro Lys Pro Lys Xle Pro
625 630 635 640

10 Lys Glu Pro Glu Glu Gly Glu Thr Pro Pro Glu Ser Ala Lys lie Gly
645 650 655

Thr Arg Lys Phe Tyr Arg Lys Lys Arg Trp Val Asp Ala Gin Leu Tyr
660 665 * 670

His Met Glu Ser Leu Arg Pro Gly Asn Arg Val Met Gly Pro Ala Val

675 680 685

He Glu Ser Asp Ala Thr Thr Phe Val Val Pro Asp Gly Phe Glu Thr
20 690 695 700

Trp Leu Asp Gly His Arg Leu Phe His Leu Arg Glu Val
705 710 715

<210> 3
<211> 168
<212> PRT

<213> Xanthobacter autotrophics Py2
<220>

<221> DOMAIN
<222> 1. .168

<223> Acetone carboxylase, gamma subunit
<400> 3

Met Ala Tyr Thr Arg Ser Lys He Val Asp Leu Val Asp Gly Lys He
1 .5 10 15

40 Asp Pro Asp Thr Leu His Gin Met Leu Ser Thr Pro Lys Asp Pro Glu
20 25 30

Arg Phe Val Thr Tyr Val Glu He Leu Gin Glu Arg Met Pro Trp Asp
35 40 45

Asp Lys He He Leu Pro Leu Gly Pro Lys Leu Phe He Val Gin Gin
50 55 60

Lys Val Ser Lys Lys Trp Thr Val Arg Cys Glu Cys Gly His Asp Phe
50 65 70 75 80

Cys Asp Trp Lys Asp Asn Trp Lys Leu Ser Ala Arg Val His Val Arg
85 90 95

55 Asp Thr Pro Gin Lys Met Glu Glu He Tyr Pro Arg Leu Met Ala Pro
100 105 110

Thr Pro Ser Trp Gin Val He Arg Glu Tyr Phe Cys Pro Glu Cys Gly
115 120 125

107

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Thr Leu His Asp Val Glu Ala Pro Thr Pro Trp Tyr Pro Val He His
130 135 140

Asp Phe Ser Pro Asp He Glu Gly Phe Tyr Gin Glu Trp Leu Gly Leu

145 150 155 160

Pro Val Pro Glu Arg Ala Asp Ala
165

<210> 4
<211> 769
<212> PRT

<213> Rhodobacter capsulatus B10
<220>

<221> DOMAIN
<222> 1. .769

<223> Acetone carboxylase, alpha subunit (Capsulapedia No. RRC02651)
<400> 4

Met Asn Ala Pro Thr Ala He Arg Gly He Val Arg Gly Gly Asp Thr
1 5 10 15

25 Leu Lys Gin His Arg Asp Gly He Met Glu Ala Ser Lys Arg Thr Gly
20 25 30

His Tyr Ala Gly Leu Lys Gin Met Glu Leu Arg Asp Ser Asp Pro He
35 40 45

Met Tyr Asn Lys Leu Phe Ser Arg Leu Arg Ala Gly Val Val Asp Ala
50 55 60

Arg Glu Thr Ala Lys Lys He Ala Ala Ser Pro He Val Glu Gin Glu
35 65 70 75 80

Gly Glu Leu Cys Phe Thr Leu Tyr Asn Ala Ala Gly Asp Ser He Leu
85 90 95

40 Thr Ser Thr Gly He He He His Val Gly Thr Met Gly Ala Ala He
100 105 110

Lys Tyr Met He Glu Asn Asp Trp Glu Ser Asn Pro Gly Val Lys Asp
115 120 125

Arg Asp He Phe Cys Asn Asn Asp Ser Leu He Gly Asn Val His Pro
130 135 140

Cys Asp He His Thr He Val Pro He Phe His Glu Gly Glu Leu He
50 145 150 155 160

Gly Trp Val Gly Gly Val Thr His Val He Asp Thr Gly Ala Val Gly
165 170 175

55 Pro Gly Ser Met Thr Thr Gly Gin Val Gin Arg Phe Gly Asp Gly Tyr
180 185 190

Ser Val Thr Cys Arg Lys Val Gly Glu Asn Asp Thr Leu Phe Arg Asp
195 200 205

108

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Trp Leu His Glu Ser Gin Arg Ser Val Arg Thr Thr Arg Tyr Trp Met
210 215 220

Leu Asp Glu Arg Thr Arg lie Ala Gly Cys His Met lie Arg Lys Leu
5 225 230 235 240

Val Ala Glu Val lie Ala Glu Glu Gly He Glu Ala Tyr Trp Lys Phe
245 250 255

10 Ala Tyr Glu Ala Val Glu His Gly Arg Leu Gly Leu Gin Asn Arg He
260 265 270

Lys Ala Met Thr He Pro Gly Lys Tyr Arg Gin Val Gly Phe Val Asp
275 280 285

Val Pro Tyr Ala His Asp Asp Val Arg Val Pro Ser Asp Phe Ala Lys
290 * 295 300

Val Asp Thr He Met His Thr Pro Ser Glu Met Thr He Arg Pro Asp
305 310 315 320

Gly Thr Trp Arg Leu Asp Phe Glu Gly Ala Ser Arg Trp Gly Trp His
325 330 335 .

25 Thr Tyr Asn Ala His Ser Val Ser Phe Thr Ser Gly He Trp Val Met
340 345 350

Met Thr Gin Thr Leu He Pro Thr Glu Met He Asn Asp Gly Ala Ala
355 360 365

Tyr Gly Thr Glu Phe Arg Leu Pro Lys Gly Thr Trp Met Asn Pro Asp
370 375 380

Asp Arg Arg Val Ala Phe Ala Tyr Ser Trp His Phe Leu Val Ser Ser
385 390 395 400

Trp Thr Ala Leu Trp Arg Gly Leu Ser Arg Ser Tyr Phe Gly Arg Gly
405 410 415

40 Tyr Leu Glu Glu Val Asn Ala Gly Asn Ala Asn Thr Ser Asn Trp Leu
420 425 430

Gin Gly Gly Gly Phe Asn Gin Tyr Asp Glu He His Ala Val Asn Ser

435 ' 440 . 445

Phe Glu Cys Ala Ala Asn Gly Val Gly Ala Ser Ala He Gly Asp Gly

450 455 ' 460

Leu Ser His Ala Ala Ala He Trp Asn Pro Glu Gly Asp Met Gly Asp
465 470 475 480

Met Glu He Trp Glu Leu Ala Glu Pro Leu Val Tyr Leu Gly Arg Gin
485 490 495

55 He Lys Ala Ser Ser Gly Gly Ala Gly Lys Tyr Arg Gly Gly Cys Gly
500 505 510

Phe Glu Ser Leu Arg Met Val Trp Asn Ala Lys Asp Trp Thr Met Phe
515 520 525

109

WO 03/039483 PCT/US02/36028

Phe Met Gly Asn Gly His lie Ser Ser Asp Trp Gly Leu Met Gly Gly
530 535 " 540

Tyr Pro Ala Ala Ser Gly Tyr Arg Phe Glu Ala His Glu Thr Gly Leu
5 545 550 555 560

Lys Glu He lie Ala Gin Gly Gly Asp lie Pro His Gly Gly Asp Thr
565 570 575

10 Asp Pro Gly Asn Pro Val Trp Asp Gly Leu Leu Lys Gly Ala Arg lie
580 585 590

Lys Arg Asp Lys Gin Ala He Thr Thr Glu Ala Met Phe Lys Asp Tyr
595 600 605

Asp Leu Tyr Leu Asn Tyr Met Arg Gly Gly Pro Gly Phe Gly Asp Pro
610 615 620

Leu Asp Arg Asp Pro Gly Ala Val Ala Ala Asp Val Asn Gly Gly Tyr
20 625 630 635 "* 640

Leu Val Glu Arg Phe Ala Gin Ser Val Tyr Gly Val Val Leu Val Lys
645 650 655

25 Gly Ala Asp Gly Leu Leu Ala Ala Asp Ala Ala Ala Thr Glu Ala Arg
660 665 670

Arg Ala Ala He Arg Lys Asp Arg Leu Ala Lys Ala Val Pro Thr Ala
675 680 685

Glu Trp Met Lys Gly Glu Arg Asp Arg He Leu Lys Lys Glu Ala Gly
690 695 700

Val His Val Gin Gin Met Phe Ala Ala Ser Phe Lys Leu Gly Pro Lys
35 705 710 715 720

Trp Glu Glu Gly Phe Arg Lys Phe Trp Asp Leu Pro He Asp Trp Arg
725 730 735

40 Leu Met Glu Ala Asp Leu Pro He Pro Ser Tyr Gly Arg Asp Tyr Ser
740 745 " 750

Met Asp Leu Ser Glu Leu Pro Asp Val Lys Thr Val Gin Phe Val Glu
755 760 765

Glu

<210> 5

50 <211> 717

<212> PRT

<213> i^hodobacter capsulatus B10

<220>
55 <221> DOMAIN
<222> 1. .717

<223> Acetone carboxylase, beta subunit (Capsulapedia No. RRC02652)
<400> 5

60 Met Pro Leu Asp Arg Glu Lys Thr Arg Ser Val Gin Val Leu Gly He
1 5 10 15

110

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Asp Ala Gly Gly Thr Met Thr Asp Thr Phe Phe Val Asp Ala Asn Gly
20 25 30

Asp Phe Val Val Gly Lys Ala Gin Ser Thr Pro Gin Asn Glu Ala Leu
35 40 45

Gly Leu Leu Glu Ser Ser Arg Glu Gly Leu Gin His Trp Gly Leu Ser
50 55 60

Leu Glu Glu Ala Leu Ser Ser lie Gin Thr Gly Val Tyr Ser Gly Thr
65 70 75 80

Ala Met Leu Asn Arg Val Val Gin Arg Lys Gly Leu Arg Cys Gly Leu
85 90 " 95

He Val Asn Ala Gly Met Glu Asp Phe His Arg Met Gly Arg Ala He
.100 105 110

20 Gin Ala Tyr Leu Gly Phe Ala Tyr Glu Asp Arg He His Leu Asn Thr
115 120 125

His Tyr Tyr Asp Glu Pro Leu Val Pro Arg His Leu Thr Arg Gly Val

130 135 140

Met Glu Arg He Asp Met Phe Gly Asp Val Val He Pro Leu Arg Glu

145 150 155 160

Glu Thr Ala Arg Gin Ala Ala Ala Glu Leu He Ala Gin Asp Val Glu
165 170 175

Gly He Val He Ser Leu Leu His Ser Tyr Lys Asn Pro Ala His Glu
180 185 190

35 . Arg Arg Val Arg Asp He Val Ala Glu Glu Leu Glu Lys Ala Gly Lys
195 200 205

Thr Thr Pro Val Phe Ala Ser Thr Asp Tyr Tyr Pro Val Arg Lys Glu

210 215 220

Thr His Arg Thr Asn Thr Thr He Leu Glu Ala Tyr Ala Ala Glu Pro

225 230 235 240

Ser Arg Gin Thr Leu Arg Lys He Thr Gly Ala Phe Lys Glu Asn Gly
45 245 250 255

Ser Arg Phe Asp Phe Arg Val Met Ala Thr His Gly Gly Thr He Ser
260 265 270

50 Trp Lys Ala Lys Glu Leu Ala Arg Thr He Val Ser Gly Pro He Gly
275 280 285

Gly Val He Gly Ala Lys Tyr Leu Gly Glu Val Leu Gly Tyr Lys Asn

290 295 300

He Ala Cys Ser Asp He Gly Gly Thr Ser Phe Asp Val Ala Leu He

305 310 315 320

Thr Gin Asn Glu Leu Thr lie Arg Asn Asp Pro Asp Met Ala Arg Leu
60 325 330 335

111

WO 03/039483 PCT/US02/36028

Val Leu Ser Leu Pro Leu Val Ala Met Asp Ser Val Gly Ala Gly Ala
340 345 350

Gly Ser Phe lie Arg Leu Asp Pro Tyr Thr Lys Ala lie Lys Leu Gly
355 360 365

Pro Asp Ser Ala Gly Tyr Arg Val Gly Val Cys Trp Ala Glu Ser Gly
370 375 380

lie Glu Thr Val Thr He Ser Asp Cys His Val He Leu Gly Tyr Leu
385 390 395 400

Asn Pro Asp Asn Phe Leu Gly Gly Gin Val Lys Leu Asp Arg Gin Arg
405 410 415

Ala Trp Asp Ala Met Lys Thr Gin He Ala Asp Pro Leu Gly Leu Ser
420 425 430

Val Glu Asp Ala Ala Ala Gly Val He Glu Leu Leu Asp Ser Asp Leu
435 440 445

Arg Asp Tyr Leu Arg Ser Met He Ser Gly Lys Gly Tyr Ser Pro Ser
450 455 460

Ser Phe Thr Cys Phe Ser Tyr Gly Gly Ala Gly Pro Val His Thr Tyr
465 470 475 480

Gly Tyr Thr Glu Gly Leu Gly Phe Glu Asp Val He Val Pro Ala Trp
485 490 495

Ala Ala Gly Phe Ser Ala Phe Gly Cys Ala Ala Ala Asp Phe Glu Tyr
500 505 510

Arg Tyr Asp Lys Ser Leu Asp Leu Asn He Ala Arg Asp Gly Ser Asp
515 520 525

Asp Leu Lys Ala His Glu Ala Arg Thr Leu Asn Asp Ala Trp His Glu
530 535 540

Leu Thr Glu Arg Val Leu Glu Glu Phe Glu Leu Asn Gly Tyr Thr Arg
545 550 555 " 560

Asp Gin Val Lys Leu Gin Pro Gly Phe Arg Met Gin Tyr Arg Gly Gin
565 570 575

Leu Asn Asp Leu Glu He Glu Ser Pro He Pro Ala Ala Lys Thr Ala
580 585 590

Ala Asp Trp Asp Lys Leu Val Ala Ala Phe Asn Asp Thr Tyr Gly Arg
595 600 605

Val Tyr Ala Ala Ser Ala Arg Ser Pro Glu Leu Gly Tyr Ser Val Thr
610 615 620

Gly Ala He Met Arg Gly Met Val Pro He Pro Lys Pro Lys He Pro
625 630 635 640

Lys Glu Pro Glu Thr Gly Ala Thr Pro Pro Glu Ala Ala Lys Leu Gly
645 650 655

112

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Thr Arg Lys Phe Tyr Arg Lys Lys Lys Trp Val Asp Ala Arg Leu Tyr
660 665 670

Arg Met Glu Lys Leu Leu Pro Gly Asn Arg lie Thr Gly Pro Ala lie
5 675 680 685

lie Glu Ser Asp Ala Thr Thr Phe Val Val Pro Asp Gly Phe Glu Thr
690 695 700

10 Trp Leu Asp Gly His Arg Leu Phe His Leu Lys Glu Val
705 710 715

. <210> 6

15 <211> 167

<212> PRT

<213> Rhodobacter capsulatus BIO

<220>
20 <221> DOMAIN
<222> 1. .167

<223> Acetone carboxylase, gamma subunit (Capsulapedia No. RRC04094)
<400> 6

25 Met Ala Tyr Thr Lys Ala Lys lie Lys Asp Leu Val Asp Gly Lys lie

1 . 5 .10 15

Asp Arg Asp Thr Leu His Thr Met Leu Ala Thr Pro Lys Asp Ala Asp
20 25 30

Arg Phe Val Met Tyr Leu Glu Val Leu Gin Asp Gin Val Pro Trp Glu
35 40 45

Asp Arg lie lie Leu Pro Leu Gly Pro Lys Leu Tyr lie Val Gin Arg
35 50 55 60

Lys Ser Asp His Lys Trp Val Val Lys Ser His Ala Gly His Glu Phe
65 70 75 80

40 Cys Asp Trp Arg Glu Asn Trp Lys Leu His Ala Val Met Arg Val Arg

85 90 95

Glu Thr Pro Glu Ala Met Glu Glu He Tyr Pro Arg Leu Met Ala Pro
100 105 110

Thr Ala Gly Trp Gin Val He Arg Glu Tyr Tyr Cys Pro Leu Ser Gly
115 120 125

Asp Leu Leu Asp Val Glu Ala Pro Thr Pro Trp Tyr Pro Val He His
50 130 135 140

Asp Phe Glu Pro Asp He Asp Ala Phe Tyr Ser Glu Trp Leu Gly Leu
145 150 155 160

55 Lys He Pro Glu Arg Ala Ala

165

<210> 7
60 <211> 14
<212> PRT

113

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<213> Xanthobacter autotrophics Py2
<220>

<221> DOMAIN
5 <222> 1. .14

<223> Secondary alcohol dehydrogenase, N-terminal tetradecapeptide
<400> 7

Met Lys Gly Leu Val Tyr Arg Gly Pro Gly Lys Lys Ala Leu
10 1 5 10

<210> 8
<211> 12
15 <212> PRT

<213> Xanthobacter avtotrophicus Py2

<220>

<221> DOMAIN
20 <222> 1. .12

<223> Secondary alcohol dehydrogenase, tryptic fragment

<220>

<221> UNSURE

25 <222> 9. .9

<223> Phe9 may be Ser9

<400> 8

Pro Val Ala Val Asp His Gly Pro Phe Pro His Lys
30 1 5 10

<210>

<211>

<212>

PRT

<213>

Xanthobacter autotrophics Py2

<220>

<221>

DOMAIN

<222>

1. .8

<223>

Secondary alcohol dehydrogenasi

<220>

<221>

UNSURE

<222>

3. .3

<223>

Leu3 may be Ile3

<400>

Gly Gly Leu Gly Val Tyr His Gin

<210>

<211>

<212>

PRT

<213>

Xanthobacter autotrophicus Py2

<220>

<221> DOMAIN
60 <222> 1. .9

<223> Secondary alcohol dehydrogenase, tryptic fragment

114

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<220>

<221> UNSURE
<222> 2. .2
5 <223> Leu2 may be Ile2

<400> 10

Ala Leu Glu Glu Val Pro His Pro Arg
1 5

<210> 11
<211> 7
<212> PRT

<213> Xanthobacter autotrophicus Py2

.c^ DOMAIN

<222> 1..7

j <223> Secondary alcohol dehydrogenase, tryptic fragment .

<400> 11

His Pro Ser Gly Asp Thr Arg
1 5

<210> 12 .

<211> 9

<212> PRT

30 <213> Xanthobacter autotrophicus Py2
<220>

<221> DOMAIN

<222> 1. .9

35 <223> Secondary alcohol dehydrogenase, tryptic fragment

<400> 12

Gly Leu Val Tyr Arg Gly Pro Gly Lys
1 5

40 .

<210> 13

<211> 7

<212> PRT

45 <213> Xanthobacter autotrophicus Py2
<220>

<221> DOMAIN

<222> 1..7

50 <223> Secondary alcohol dehydrogenase, tryptic fragment
<220>

<221> UNSURE

<222> 3. .3

55 <223> Ile3 may be Leu3

<400> 13

His Gin lie Ala Ser Ser Arg
1 5

115

WO 03/039483

PCT/US02/36028

<210> 14
<211> 6
<212> PRT

<213> Xanthobacter autotrophics Py2

<220>

<221> DOMAIN
<222> 1. .6

<223> Secondary alcohol dehydrogenase, fragment

<400> 14

Leu Asp Asn Val Pro Glu
1 5

<210> 15
<211> 6
<212> PRT

<213> Xanthobacter autotrophics Py2

<220>

<221> DOMAIN
<222> 1. .6

<223> Secondary alcohol dehydrogenase, fragment

<400> 15

Phe Asp Gin Arg Gin Pro

1.5

<210> 16
<211> 8
<212> PRT

<213> Xanthobacter au totrophicus Py2

<220>

<221> DOMAIN
<222> 1. .8

<223> Secondary alcohol dehydrogenase, fragment

<400> 16

Gly Ala Gly Arg He He Ala Val
1 5

<210> 17
<211> 7
<212> PRT

<213> Xanthobacter a u totrophicus Py2

<220>

<221> DOMAIN
<222> 1. .7

<223> Secondary alcohol dehydrogenase, fragment

<<300> 17

Gin Val Glu Pro Leu Met Ser
5

60 <210> 18
<211> 9

116

WO 03/039483

PCT/US02/36028

<212> PRT

<213> Xanthobacter autotrophics Py2

. <220>
5 <221> DOMAIN
<222> 1..6

<223> Secondary alcohol dehydrogenase, fragment
<400> 18

10 Phe Phe Ala Asp lie lie Glu Ala Ala
1 5

<210> 19
15 <211> 6

<212> PRT

<213> Xanthobacter avtotrophicus Py2

<220>
20 <221> DOMAIN
<222> 1. .6

<223> Secondary alcohol dehydrogenase, fragment

<400> 19
25 Asp Thr Val Thr Thr His
1 5

117

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